Effect of Knockdown of Genes Involved in the RNAi Pathway ... · Effect of Knockdown of Genes Involved in the RNAi Pathway on Root-knot Nematodes This thesis is presented to Murdoch

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Effect of Knockdown of Genes Involved in the RNAi

Pathway on Root-knot Nematodes

This thesis is presented to Murdoch University for the degree of

Doctor of Philosophy

By

Sadia Iqbal

M.Sc (Hons.)

Western Australian State Agriculture Biotechnology Centre

School of Veterinary and Life Sciences

Murdoch University

Perth, Western Australia

2015

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Declaration

I declare that this thesis is my own account of my research and contains as its main content,

work which has not been previously submitted for a degree at any tertiary educational

institution.

_____________

Sadia Iqbal

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Abstract

Plant parasitic nematodes are economically important crop pests: of these root-knot nematodes

(Meloidogyne spp.) have the widest host range and are recognised as the most significant

nematode pests of crop plants worldwide. Current control methods have serious limitations:

natural resistance genes, chemicals and cultural practices are often not effective or are too

expensive for large scale application. An alternative control strategy chosen here was to target

and silence a conserved mechanism (RNA interference – RNAi) in the nematodes via transgenic

plants. The RNAi pathway itself was studied for its potential to provide new gene targets for

nematode control, with potentially wider applications to control of other plant nematodes.

In this study the overall aim was to undertake in silico identification of genes and ‘effectors’ of

the RNAi pathway of root-knot nematodes, to use RNAi to silence some of these target genes,

and to determine the effects on their parasitic success after both in vitro and in planta RNAi

treatments. Twenty-seven genes were identified in the RNAi pathways of M. incognita, and

selected for further study. In vitro RNAi experiments (‘soaking’ of J2 nematodes in dsRNA

homologous to the target gene) to down-regulate expression of the 27 selected genes caused

significant effects on the infectivity and development of the nematodes when they were used to

infect susceptible tomato plants. Up to a 90% reduction in infection was observed for dcr-1

targeted nematodes. Down-regulation of effectors of the miRNA pathway (drsh-1, pash-1, alg-

1, xpo-1, xpo-2) and dicer complex (drh-1, drh-3) had the greatest effect on nematode viability

and/or development.

Seven of the cloned genes (dcr-1, drh-3, vig-1, mut-7, drsh-1, pash-1, rha-1) were chosen after

in vitro screening for in planta analysis, and hairpin constructs for each were successfully

transformed into A. thaliana plants. Challenge of the heterozygous T2 transgenic plants with M.

incognita J2s exhibited significant reductions in infection parameters: 31 transgenic events of

the 7 genes showed a reduction of infection of 50% or more when compared with controls, and

the greatest reduction was 89%, for plants targeting drsh-1of M. incognita.

Another in vitro RNAi experiment targeting 7 different regions of the same gene, dcr-1, was

conducted to evaluate gene silencing in relation to different regions of the same target. The

results showed that there was variable target expression and RNAi effects depending on the

target region used, with higher impact for sequences near the 5′ end of the targeted transcript.

Targeting dcr-1 resulted in reduced nematode infection and reproduction: abnormal nematode

development was also observed.

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This project provides new information on genes involved in small RNA pathways of M.

incognita, resulting in identification of novel targets for its control by gene silencing

technology. It also provides additional data to improve design of more effective RNAi triggers

related to the target region chosen. The in planta RNAi results generated provide further

evidence of the potential of RNAi as a nematode control strategy based on results using the

model plant A. thaliana: it is likely that these results are translatable to protect crop plants from

nematode attack in the future.

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List of Conferences and Publications

Stephen J. Wylie, Chao Zhang, Vicki Long, Marilyn J. Roossinck, Koh S-H, Michael G.K.

Jones, Sadia Iqbal, Hua Li. 2015. Differential responses to virus challenge of laboratory and

wild accessions of Australian species of Nicotiana, and comparative analysis of RDR1 gene

sequences. PloS one, 10(3), e0121787

Sadia Iqbal, John Fosu-Nyarko and Michael G.K. Jones. 2015. Genomes of parasitic

nematodes (Meloidogyne hapla, Meloidogyne incognita, Ascaris suum and Brugia malayi) have

a reduced complement of small RNA interference pathway genes: knockdown of some reduces

host infectivity of M. incognita. Submitted in Functional & Integrative Genomics.

Sadia Iqbal, John Fosu-Nyarko and Michael G.K. Jones. 2012. Silencing the effectors of RNA

silencing. 31st International Symposium of the European society of nematologists. Adana,

Turkey.

Sadia Iqbal, John Fosu-Nyarko and Michael G. K. Jones. 2012. Silencing the effectors of RNA

silencing. Combined Biological Sciences Meeting. Perth, Western Australia.

Sadia Iqbal, John Fosu-Nyarko and Michael G. K. Jones. 2015. RNA silencing of RNAi

effectors reduces root-knot nematode infection. Australasian Plant Pathology Society

Conference. Perth, WA.

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Table of Contents

Declaration ..................................................................................................................................... I

Abstract ......................................................................................................................................... II

List of Conferences and Publications .......................................................................................... IV

Table of Contents ........................................................................................................................ VI

Abbreviations ................................................................................................................................ X

Acknowledgement ..................................................................................................................... XII

Chapter 1 Literature review .......................................................................................................1

1.1 Introduction to nematodes ........................................................................................................2

1.1.1 Plant parasitic nematodes (PPNs) ..................................................................................3

1.1.2 Root-knot nematodes (RKNs) ........................................................................................4

1.1.3 Life cycle of RKNs ........................................................................................................5

1.1.4 Methods of PPN control .................................................................................................8

1.2 RNAi or gene silencing ..........................................................................................................10

1.2.1 RNAi mechanism .........................................................................................................12

1.3 Components of siRNA and miRNA pathways of eukaryotes ................................................15

1.3.1 Transport proteins ........................................................................................................15

1.3.2 Dicer complex ..............................................................................................................16

1.3.3 RNA-induced silencing complex (RISC) ....................................................................17

1.3.4 RNAi amplification machinery ....................................................................................19

1.3.5 RNAi inhibitors ............................................................................................................19

1.3.6 Nuclear RNAi ..............................................................................................................20

1.3.7 Argonautes ...................................................................................................................22

1.4 Applications of RNAi ............................................................................................................25

1.4.1 Development of in vitro RNAi in nematodes ..............................................................26

1.4.2 In vitro RNAi of PPN genes ........................................................................................27

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1.4.3 Application of virus-induced gene silencing to PPN research .....................................29

1.4.4 The potential for PPN control using RNAi ..................................................................30

1.5 Aims and objectives of this project ........................................................................................33

Chapter 2: Genome level identification and comparison of effectors of RNAi pathway of

the parasitic nematodes Meloidogyne hapla, Meloidogyne incognita, Ascaris suum and

Brugia malayi. .............................................................................................................................36

2.1 Abstract ..................................................................................................................................37

2.2 Introduction ............................................................................................................................37

2.3 Materials and methods ...........................................................................................................38

2.3.1 Identification of effectors of RNAi of C. elegans .........................................................38

2.3.2 Identification of genomic contigs of parasitic nematodes mapped to RNAi effectors of

C. elegans. ..............................................................................................................................39

2.3.3 In silico functional analysis of putative effectors of the parasitic nematodes ...............39

2.3.4 Phylogenetic analyses ...................................................................................................40

2.4 Results ....................................................................................................................................40

2.4.1 Genomic contigs and ESTs of M. incognita, M. hapla, A. suum and B. malayi with

homologies to effectors of C. elegans.....................................................................................40

2.4.2 Small RNA transport proteins .......................................................................................41

2.4.3 The Dicer and associated genes ....................................................................................43

2.4.4 RNA-induced silencing complex (RISC)......................................................................45

2.4.5 RNAi amplification .......................................................................................................46

2.4.6 RNAi inhibitors .............................................................................................................47

2.4.7 Nuclear RNAi effectors ................................................................................................50

2.4.8 Argonautes ....................................................................................................................50

2.5 Discussion ..............................................................................................................................52

Chapter 3: Identification of target genes from among sRNA pathway effectors of M.

incognita for nematode control via in vitro RNAi ...................................................................57

3.1 Abstract ..................................................................................................................................58

3.2 Introduction ............................................................................................................................59


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3.3.1 Primer design ...............................................................................................................60

3.3.2 RNA extraction ............................................................................................................61

3.3.3 Amplification of target genes .......................................................................................62

3.3.4 Cloning of amplicons into RNAi vector ......................................................................62

3.3.5 Confirmation of sequences cloned into vectors ...........................................................64

3.3.6 Synthesis of dsRNA .....................................................................................................64

3.3.7 In vitro feeding of dsRNA to nematodes .....................................................................64

3.3.8 Observation of RNAi phenotypes ................................................................................65

3.3.9 Assessment of nematode infectivity and development ................................................65

3.3.10 Gene expression of target genes .................................................................................66

3.3.11 Statistical analysis ......................................................................................................66

3.4 Results ....................................................................................................................................66

3.4.1 Phenotypic effects of in vitro RNAi of target genes ....................................................66

3.4.2 RNAi of target genes of nematodes reduces host infection .........................................69

3.4.3 RNAi effects on nematode development .....................................................................72

3.4.4 Transcript abundance after in vitro feeding .................................................................75

3.5 Discussion ..............................................................................................................................77

Chapter 4: Host-induced gene silencing of RNAi effectors confers resistance against

Meloidogyne incognita and affects development......................................................................81

4.1 Abstract ..................................................................................................................................82

4.2 Introduction ............................................................................................................................82


4.3.1 Cloning of hairpin expression cassettes ........................................................................83

4.3.2 Cloning into the binary vector pART27........................................................................85

4.3.3 Agrobacterium tumefaciens transformation ..................................................................86

4.3.4 Plant transformation ......................................................................................................86

4.3.5 Screening for Transgenic (T1) plants:...........................................................................87

4.3.6 Screening and challenge of T2 plants ...........................................................................88

4.3.7 Nematode collection for infection .................................................................................88

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4.3.8 Nematode infection of transgenic plants .......................................................................88

4.3.9 Nematode development after infection .........................................................................89

4.3.10 Statistical analyses ......................................................................................................89

4.3.11 Confirmation of T-DNA insertion ..............................................................................89

4.3.12 Confirmation of transcription of nematode silencing signals .....................................90

4.4 Results ....................................................................................................................................90

4.4.1 Analysis of transgenic plants .......................................................................................90

4.4.2 Analysis of nematode infection ....................................................................................92

4.4.3 Female morphology .....................................................................................................95

4.4.4 T-DNA insertion and dsRNA transcription .................................................................96

4.5 Discussion ..............................................................................................................................97

Chapter 5: The effects of RNAi treaments with different regions of the Dicer-like gene on

the viability, parasitism and reproduction of M. incognita ..................................................100

5.1 Abstract ................................................................................................................................101

5.2 Introduction ..........................................................................................................................101

5.3 Materials and Methods .........................................................................................................102

5.3.1 Sequence analysis and Primer design ........................................................................102

5.3.2 Cloning and synthesis of dsRNAs .............................................................................103

5.3.3 In situ hybridisation ...................................................................................................103

5.3.4 dsRNA soaking ..........................................................................................................104

5.3.5 Phenotype, RNA extraction and plant infection ........................................................104

5.3.6 Analysis of infection ..................................................................................................105

5.3.7 Quantification of gene knockdown ............................................................................105

5.3.8 Statistical analysis ......................................................................................................106

5.4 Results ..................................................................................................................................107

5.4.1 Expression pattern of dcr-1 in J2 M. incognita ..........................................................107

5.4.2 RNAi phenotype for dcr-1 domains ...........................................................................107

5.4.3 Nematode infection after dcr-1 RNAi and targeted region ........................................108

5.4.4 Differential effects of target region on reproduction .................................................109

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5.4.5 Nematode development is affected by treatment with dcr-1 RNAi ...........................110

5.4.6 Target quantification ..................................................................................................111

5.4.7 RNAi of dcr-1 affects other RNAi effectors ..............................................................112

5.4.8 Target and trigger properties affecting RNAi efficiency ...........................................114

5.5 Discussion ............................................................................................................................115

Chapter 6: General Discussion ...............................................................................................117

6.1 Overview ..............................................................................................................................118

6.2 Effectors of small RNA pathways of RKN ..........................................................................119

6.3 In vitro RNAi as a functional analysis tool for parasitic nematodes ....................................120

6.4 HIGS for RKN control .........................................................................................................121

6.5 The target region of a gene affects RNAi effectiveness ......................................................122

6.6 Future directions ..................................................................................................................123

6.7 Conclusions ..........................................................................................................................124

APPENDIX ................................................................................................................................126

References ..................................................................................................................................131

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Abbreviations

APN Animal parasitic nematode

As Ascaris suum

ATP Adenosine tri-phosphate

Bm Brugia malayi

bp Base-pair

Ce Caenorhabditis elegans

CN Cyst nematode

Ct Cycle threshold

Cv Cultivar

DCL Dicer-like

DNA Deoxyribonucleic acid

dsRNA Double stranded RNA

Endo-RNAi Endogenous RNA interference

Exo-RNAi Exogenous RNA interference

GFP Green fluorescent protein

HIGS Host-induced gene silencing

Hp Hairpin

J1 First-stage juvenile

J2 Second-stage juvenile

J3 Third-stage juvenile

J4 Fourth-stage juvenile

Mh Meloidogyne hapla

Mi Meloidogyne incognita

miRISC micro RNA induced silencing complex

miRNA MicroRNA

mRNA Messenger RNA

nt Nucleotide

P bodies Processing bodies

PCR Polymerase chain reaction

Pi-RNA PIWI interacting RNA

PPN Plant parasitic nematode

Pre-mRNA Precursor messenger RNA

Pre-miRNA Precursor microRNA

Pri-miRNA Primary microRNA

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PTGS Post transcriptional gene silencing

qRT-PCR quantitative real-time polymerase chain reaction

RdRp RNA-directed RNA polymerase

RISC RNA-induced silencing complex

RKN Root-knot nematode

RNA Ribonucleic acid

RNAi RNA interference

rRNA ribosomal RNA

siRISC short interfering RNA induced silencing complex

siRNA Short interfering RNA

SNase Staphylococcol nuclease

sRNA Small RNA

VIGS Virus induced gene silencing

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Acknowledgement

Firstly, I would like to thank the Endeavour Awards Australia for giving me a Postgraduate

Research Scholarship to do this Ph.D. I would like to express my gratitude to my supervisors,

Professor Michael G.K. Jones and Dr John Fosu-Nyarko for all the time, guidance and

motivation they have given me. Their invaluable support and encouragement helped me in

staying positive throughout.

My sincere thanks go to everyone in SABC for their help. Special thanks to Frances Brigg for

all the sequencing help and Gordon Thompson for help with microscopy, Dave Berryman, Bee

Lay Addis and Karen Olkowski for all the support they have provided towards completing this

thesis. I would like to thank the experienced researchers Dr. Steve Wylie, Dr. Reetinder Kaur,

Dr. Vaughan Agrez, Meenu Singh and Dr. Leila Eshraghi for sharing their knowledge and

experiences which helped me in completing this project.

A big thankyou to the Ph.D students, Jo-Anne Tan, Jamie Ong, Shu Hui Koh and Elvina Lee for

their support, valuable discussions and troubleshooting advice which helped me at many steps

during this project and gave me friends for life. I would also like to thank colleagues in the Plant

Biotechnology Research Group (Fareeha Naz, Farhana Begum, Harshini Herath, Malathy

Rathinasamy, Sharmin Rahman Silvee and Vineeta Bilgi) for all the cheerful get togethers and

time we spent in the office.

And most importantly, thanks to my parents and siblings for their unconditional love and

support in my entire life. Special thanks to my husband for being there for me and helping me in

every way possible and my friends in Pakistan who supported me and became my punching

bags whenever needed during the course of this Ph.D.

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Dedicated to

MY PARENTS

who have supported me in my every decision

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Chapter 1

Literature Review

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1.1 Introduction to nematodes

Nematodes are unsegmented roundworms which have a simple cylindrical body, generally 250

µm to 12 mm in length and 15-35 µm in width (Lambert and Bekal 2002). The phylum

Nematoda is a diverse and abundant group of metazoans with 25,000 species described so far

out of an estimated one million existing species (Bongers and Bongers 1998; Lambshead and

Boucher 2003). New sequencing technologies have made it feasible to gain insight into the

evolution, diversity and relationship of organisms, and this also applies to nematodes. Their

ecological diversity ranges from soil inhabiting nematodes to marine and terrestrial habitats, and

even to extreme environments such as the polar regions. The life span of nematodes ranges from

a few days for Caenorhabditis elegans to 15 years for the human parasitic hookworm

Necatoriasis americanus (Palmer 1955). The free-living bacteriovorous nematode C. elegans

was the first multicellular organism to have its genome sequenced. It has a genome of 100 Mbp

and ~20,000 annotated genes (C. elegans Sequencing Consortium 1998). Like other animals, C.

elegans has structures that include a digestive system, nervous system, muscles and ‘skin’,

called the cuticle. It has been used as a model organism to study various aspects of animal

biology such as the organisation of a simple nervous system, and for functional genetics,

because of the simple biology, short life span and ease of producing genetic mutants.

Based on the phylogenetic analysis of the small subunit ribosomal RNA (rRNA) gene of

nematodes, the phylum Nematoda is divided into five clades (Blaxter et al. 1998). Free-living

nematodes in soil play an important role in nutrient turnover by making them available for plant

absorption. This makes soil inhabiting free-living nematodes important indicators of soil health

(Neher 2000). Parasitic nematodes cause economic and health problems worldwide. Animal

parasitic nematodes include disease-causing worms of humans and animals parasitising both

invertebrates and vertebrates, including domestic animals, reducing their health and leading to

economic losses for production animals. The phylogenetic relationship of nematodes is

indicated in Figure 1.1.

The ecology of soil includes study of microscopic organisms like bacteria, fungi, protozoans,

with the largest animal component being nematodes. Plant parasitic nematodes (PPNs), which

have also been termed “The Invisible Enemy” are major pests of agricultural crops, and can be

found infesting crops from temperate to tropical environments: so far 4,100 species have been

described (Decraemer and Hunt 2006; Perry and Moens 2011).

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FIGURE OMITTED

Figure 1.1: Phylogenetic relationship of nematodes based on the small subunit ribosomal RNA

gene. The five nematode clades and mode of feeding are indicated with parasitic species present

in all clades. Some important species are also indicated (Blaxter 2011).

1.1.1 Plant parasitic nematodes (PPNs)

In addition to free-living nematodes, soil inhabiting nematodes include PPNs that feed on and

parasitise plants (Decraemer and Hunt, 2006). They can infest almost all parts of plants

depending on the species: some can infest trees (Bursaphelenchus spp.) and leaves

(Aphelenchoides spp.). However, the most common nematode infestations occur in the below

ground parts of plants particularly in roots, tubers and bulbs. The life styles of PPNs can be

ecto-parasitic or endo-parasitic as defined by their feeding habit (outside or within the root

respectively), and migratory or sedentary depending on their mobility. They can have multiple

hosts (polyphagous) or may feed specifically on one or a few related species. Crops can be

infested with one or many different species of PPNs. In tropical and semi-tropical environments

it is quite clear that some form of nematode management should be undertaken to enhance

vegetable crop production, because of the pressures of nematode infestation (Sikora and

Fernandez 2005).

Migratory endoparasitic nematodes navigate their way from cell-to-cell damaging cell walls and

acquiring nutrients without inducing permanent feeding sites and most can move into and out of

the plant (Hussey and Grundler 1998). Examples of migratory nematodes include the burrowing

nematode (Radopholus spp.), root lesion nematodes (Pratylenchus spp.), pine wilt nematodes

(Bursaphelenchus spp.) and the foliar nematode (Aphelenchoides spp.). Entry into host tissues is

4

achieved using the mouth stylet and possibly by secretion of various proteins which may

include cell wall degrading enzymes.

In contrast, sedentary endoparasitic nematodes invade plant roots and modify host cells to

support long term feeding: once feeding cells are induced the nematode looses the ability to

move and feeds from these cells for the rest of its life-cycle. Examples of sedentary

endoparasitic nematodes include root-knot nematodes (Meloidogyne spp.) and cyst nematodes

(Globodera spp. and Heterodera spp). Unlike, migratory nematodes, whose feeding kill the

cells, those modified into feeding structures by sedentary nematodes remain alive and

metabolically active until the nematodes complete their life cycles. The life cycle of root-knot

nematodes (RKNs) is discussed in more detail in Section 1.1.3. Economically, the most

damaging PPNs are Meloidogyne species, commonly known as RKNs, because infested roots

develop galls or knots as a result of the feeding structures created by these nematodes (Figure

1.2).

The damage caused to world agriculture by PPNs is difficult to quantify accurately, but has

been estimated to be $125 billion per annum by Chitwood (2003), whereas the damage caused

by Meloidogyne spp. alone is estimated at about $100 billion per annum (Bird et al. 2009). This

contradiction in estimates of losses caused by PPN infestation needs to be updated using new

survey data to provide an accurate figure of the losses they cause to world agricultural

production. One aspect that confuses assessment of PPN damage is that the above ground

symptoms after nematode infection are often similar to those of plant nutrient deficiency and the

extent of damage may remain under-estimated and misdiagnosed (Hunt et al. 2005).

1.1.2 Root-knot nematodes (RKNs)

Initially reported as a root-knot disease on cucumber roots in 1855, the RKN was first described

in 1879 by Cornu. RKNs (Meloidogyne spp.) are sedentary plant endoparasites which have the

Figure 1.2: Galls resulting from infection of tomato roots by the root-knot nematode M.

incognita.

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unique ability to modify host cells into giant cells which provide nutrients for associated

nematodes’ feeding (Jones 1981; Gheysen and Fenoll 2002). Ranked as number one in

importance out of the top 10 PPNs of the world, RKN species infest plants in all temperate and

tropical cropping areas, with 98 species described so far (Jones et al. 2013). With more than

5,500 species of plants identified as hosts, RKNs infect almost every crop grown, and hence are

considered one of the most damaging plant pests (Sasser 1980; Trudgill and Blok 2001).

Crop rotation or leaving the land fallow results in reduced nematode populations but many weed

species are hosts of RKNs, and this helps them to survive and even multiply in the absence of a

cultivated crop. The population dynamics of RKNs have been studied extensively, and threshold

levels depend on the nematode species and their virulence for infecting a particular host. On

average, any soil having 0.5-2 J2s (second-stage juveniles) per gram of soil is unlikely to

provide an optimum crop yield if susceptible. Damage thresholds range from 0.01 eggs/cm3 of

soil for M. artiellia on chickpea to 10 eggs/cm3 of soil for M. incognita on corn (Di Vito et al.

1980; Di Vito and Greco 1988).

1.1.3 Life cycle of RKNs

The life cycle of Meloidogyne spp. varies from three weeks to months depending on factors

such as species, temperature, moisture and plant host (Taylor and Sasser 1978). The normal life

cycle allows them to multiply several times during one cropping season, resulting in severe

damage to host crops. Embryos of RKNs develop as first-stage juveniles (J1s) through

embryogenesis, and moult once inside the egg to become second-stage juveniles (J2s): these

hatch from the eggs when developmental and environmental conditions are favourable (Moens

et al. 2009). In contrast to cyst nematodes (CNs) which hatch in response to root exudates or

ions such as zinc chloride, RKN J2s generally hatch in water without the aid of root exudates or

added chemicals. However, their hatching is dependent on temperature. The delayed

embryogenesis and subsequent hatching of nematodes in response to unfavourable conditions

has been termed as ‘diapause’, and occurs in some species of RKNs (de Guiran 1979; Antoniou

and Evans 1987).

Active J2s locate growing root tips by sensing gradients of compounds from root exudates and

exploring root epidermal cells to find suitable points for entry into root tissues (Curtis et al.

2009). The attraction depends on the plant species, root epidermal cells and on chemotaxis in

response to carbon dioxide gradients (Robinson 1995; Zhao et al. 2000; Rodger et al. 2003). J2s

usually penetrate the host plant at the zone of root elongation (Figure 1.3), creating access by

using their needle-like stylet, accompanied by secretion of proteins from their pharyngeal gland

cells: these may include cell wall degrading enzymes. The latter are thought to soften affected

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cell walls and so enable the nematode to penetrate the root and migrate intercellularly, that is in

the apoplastic (cell wall) space of root tissues (Hussey and Grundler 1998; Abad et al. 2003).

The proteins secreted into the host by the infective J2s are selective in nature and appear to

embody different functions, for example degrading or modifying specific host cell wall

components to allow migration within the root, suppression of host defences and modifying

endogenous hormone levels and plant signalling pathways (Smant and Jones 2011; Haegeman

et al. 2012). Other nematode secretions targeted to cell contents are required to manipulate and

re-program cell development by inducing the formation of giant cells from immature vascular

elements (Jones and Payne 1978; Davis et al. 2004; Davis and Mitchum 2005).

FIGURE OMITTED

The secreted components which alter host cell development, down-regulate host defences and

are otherwise vital for successful parasitism are generally known as “effectors”. The number of

giant cells induced at the permanent feeding site can vary, usually in the order of five to seven

cells but can range from two to twelve. Following secretions by the nematode, individual cells

de-differentiate and enlarge by a process of repeated nuclear divisions without cytokinesis, to

become multinucleate without a prominent central vacuole (Jones and Payne 1978; Jones 1981;

Figure 1.3: Infection cycle of a root-knot nematode (N), illustrating the different stages of the

life cycle during infestation. Migration of the J2 nematodes in the cortex is followed by

turning and migration inside the endodermis (En): secretions into pro-vascular cells at the

feeding site lead to the modification of these cells to form giant cells (GCs), from which the

nematode feeds. Following moulting to reach the adult stage, eggs are secreted in a gelatinous

egg mass at the surface of the root. Other abbreviations: xylem (Xy), phloem (Ph), giant cells

(GC) (Bartlem et al. 2013).

7

Wiggers et al. 1990; Sijmons et al. 1994; Gheysen and Mitchum 2011). In principle, the

numbers of nuclei in developing giant cells should follow a geometric progression, but some

nuclei become polyploid, for example, 80 nuclei were counted in a single giant cell by Wiggers

et al. (1990), with each nucleus being polyploid. Division and expansion of adjacent cells of the

pericycle and cortex result in the formation of a gall or ‘knot’ surrounding the developing giant

cells, and this enables the giant cells to expand in size without damaging the surrounding cells

(Jones and Payne 1978; Jones 1981; Bird et al. 2009). This altered plant morphology is also

accompanied by up-regulation and down-regulation of expression of many genes in giant cells

compared to ‘normal’ plant cells, although deciding exactly the cell types with which to make

such comparisons is difficult (Jones 1981; Goverse et al. 2000; Gheysen and Fenoll 2002; Wang

et al. 2003; Baum et al. 2007). These changes in gene expression and metabolism enable the

giant cells to meet the demands of the feeding nematode throughout its life cycle. In a

susceptible host, the metabolism of the giant cells must match the nutrient demands of the

feeding nematode. This is reflected in the ‘transfer cell’ nature of giant cells, characterised by

the development of extensive wall ingrowths which increase the area of the cell membrane for

solute uptake from the apoplast to the symplast, and the many mitochondria required to supply

the energy needed for metabolic synthesis and solute transport – the associated nematode acts as

a nutrient sink, and the giant cell has to respond to removal of nutrients and maintain

homoestasis (Jones 1981; Caillaud et al. 2008). Galls induced by different species of

Meloidogyne differ in form and size, for example, more roots tend to develop from galls of M.

hapla than from galls induced by other RKNs.

In terms of nematode development, once J2s have initiated giant cells, they convert their lipid

reserves to glycogen and lose their ability to move. As feeding progresses, they undergo two

non-feeding moults at the feeding site i.e. J2 to J3 followed by J3 to J4, to become feeding

pyroform adult females (Hussey and Grundler 1998). During the non-feeding moults, they

survive on stored glycogen reserves. Reproduction is mostly by mitotic parthenogenesis

although males exist in all known species. M. hapla can reproduce by sexual reproduction and

has been used as a model system for genetic studies of RKNs (Liu and Williamson 2006; Blok

et al. 2008). In favourable conditions, several hundred eggs are laid into a gelatinous matrix by

each female on the surface of galled roots (Karssen and Moens 2006). The gelatinous matrix is

composed of glycoprotein which is transparent initially but turns dark brown with age. This

matrix protects the eggs from desiccation and has antimicrobial properties (Orion and Kritzman

1991).

The altered physiology of the vascular system of infected plants disrupts water and nutrient

uptake and translocation, resulting in loss of vigour and reduced crop yields and quality.

8

Severely infected plants become susceptible to secondary infections by soil fungi, bacteria and

viruses (Manzanilla-Lopez et al. 2004). This accelerates decay and death of the infected root

tissues.

1.1.4 Methods of PPN control

The overall aim of effective pest control is to use minimum resources to gain maximum benefit.

RKNs have wide host ranges, for example they have been reported to affect 138 weed species

which can be hosts and prevent disrupting infection cycles (Rich et al. 2009). The wide host

range of RKNs means that it is virtually impossible to eliminate them completely from a

cropping area. Methods of control include crop rotation, clean cultivation and chemicals but

application of nematicides like aldicarb, oxamyl and methyl bromide are regarded as

environmentally hazardous. Restrictions in their use imposed by governments have resulted in

the limitation or complete cessation of their use. Phytochemicals have also been evaluated as a

control strategy for PPNs, for example polythienyls from marigold species may be effective

against RKNs and root lesion nematodes (Ploeg 1999; Ploeg 2002; Ball-Coelho et al. 2003).

Species of the family Asteraceae when used as soil amendments or essential oil have been

reported to reduce reproduction of M. artiellia by up to 95.9% (Pérez et al. 2003). Lauric acid

from crown daisy repels M. incognita J2s and disrupts Mi flp18 expression, ultimately reducing

parasitism (Dong et al. 2014). Intercropping of such plants has been used to control RKNs, but

most natural compounds are too expensive to synthesise, unstable or otherwise not economical

or effective to use as control agents on a large scale.

Other approaches to PPN control include studying antagonistic compounds that can block

chemo-sensation employed by nematodes to locate host roots, but such effects are short-lived

(Perry 1994; Fioretti et al. 2002). Nitrogenous salts and ammonia released from decaying

organic matter also repel PPNs like M. incognita (Castro et al. 1991). Obligate parasites of

nematodes, such as Bacillus penetrans and Pasteuria penetrans have also been studied as

biological control agents. For RKNs, presence of Pasteuria penetrans in soil affects females

developing in the roots and reduces the production of egg masses (Bird and Brisbane 1988;

Davies et al. 2001; Davies et al. 2008). Arbuscular mycorrhizal fungi have also been studied as

potential biocontrol agents because their exudates can paralyse M. incognita J2s and reduce

their penetration by 32% when associated with tomato (Vos et al. 2012a; Vos et al. 2012b). The

non-pathogenic endophyte Fusarium oxysporum when inoculated on tomato plants induced

resistance to M. incognita, and resulted in a 45% reduction in nematode penetration (Dababat

and Sikora 2007). Some recently developed biological nematicides are available commercially,

but application in large fields and conditions which favour nematodes but not the biological

9

agent pose a problem for their effective use. Nevertheless, on a smaller horticultural scale some

can be effective.

It has been argued that the most sustainable method to control PPNs is the use of resistant plants

(Starr et al. 2002). Natural resistance in plants against sedentary PPNs is present in both dicot

and monocot plants. Various PPN resistance genes have been studied, including resistance to

RKNs, as presented in Table 1.1. However, most of these resistance genes are not broad

spectrum and are not available for all commercial crops. For example, the Mi gene of tomato

confers resistance against M. incognita and M. hapla, but is not effective against M. javanica -

virulent strains of nematodes can infect these resistant cultivars having Mi gene (Tzortzakakis

and Gowen 1996; Tzortzakakis et al. 1999; Jacquet et al. 2005; Castagnone-Sereno 2002).

Interestingly, this gene also confers resistance to some sucking insect pests (e.g. aphids and

whiteflies) as well (Rossi et al. 1998; Nombela et al. 2003). Apart from resistance genes against

RKNs, more than 100 soybean cyst nematode resistance sources have been studied, but the

genetic base of resistance is narrow and nematode populations are variable, which often allows

them to overcome natural resistance genes (Niblack et al. 2002; Shannon et al. 2004).

Nevertheless, there are natural resistance genes, often derived from related wild species, which

have been introduced through conventional breeding methods into crops to protect them against

nematodes, which have been used commercially. Examples include, potato cultivars with the H1

resistance gene derived from Solanum andigena which has been durable and effective against G.

rostochiensis (Starr et al. 2002).

Table 1.1: Resistance genes against RKNs identifed in various crops.

Gene RKN species Crop Reference

RMc1-hou M. chitwoodi

Potato

Draaistra 2006

RMc1-fen M. chitwoodi/ M. fallax

RMf-chc M. fallax

RMh-chcA,

RMh-chcB M. hapla

RMh-tar M. hapla

Mi-3 M.incognita/ M. javanica Tomato

Yaghoobi et al. 2005

Mi-1 M. spp. Vos et al. 1998

Mi-9 M. incognita/M. javanica/

M. arenaria

Veremis and Roberts

2000

Mae M. arenaria Peanut Garcia et al. 1996

Mag M. arenaria Peanut Garcia et al. 1996

Me3 M. incognita/M. javanica/

M. arenaria/M. hapla Pepper

Djian-Caporalino et al.

2001

Me4 M. arenaria

Me7 M. incognita Djian-Caporalino et al.

2007 Mech1 M. chitwoodi

10

Mech2 M. chitwoodi Berthou et al. 2003

However, transferring such resistance genes via transgenesis to related or unrelated species has

not worked well because downstream components that are required for internal signalling to

initiate the resistance response may not be compatible (Williamson 1999; Tai et al. 1999).

Another limitation of natural resistance genes is that they operate in a ‘gene-for-gene’ manner,

in which the resistance is based on recognition of a specific pathogen component or effector

(Gleason et al. 2008). It is therefore race specific. A pathogen race which lacks the recognised

compound will still be virulent, and so there is a constant battle between host and pathogen, in

which virulent resistance breaking pathotypes require a host mutation to recognise the pathogen

and initiate resistance responses. An exciting alternative approach to confer broader pest

resistance is based on ‘RNA interference’ (RNAi) or gene silencing technology, which uses a

conserved gene regulation mechanism present in eukaryotic cells to silence vital genes in the

pest, and so confer host resistance. This approach falls in the general category of ‘Host-Induced

Gene Silencing’ (HIGS). This is the strategy of nematode control studied in this thesis.

1.2 RNAi or gene silencing

RNAi is a highly conserved mechanism in eukaryotes which involves degradation of target

mRNAs using sequence specific small RNAs, resulting in a reduction of expression or ‘gene

knockdown’. This silencing is achieved via small RNA (sRNA) pathways which use both

exogenous and endogenous short interfering RNA (siRNA) and microRNA (miRNA) pathways.

These pathways (which are described in more detail later), interact to form a well balanced gene

regulation system (Lee et al. 2006).

Gene silencing in nematodes was first reported in C. elegans (Fire et al. 1998), and resulted in

the award of the Nobel Prize for Medicine in 2006 for Andrew Fire and Craig C. Mello. In

parallel studies on plants, this phenomenon was described in an attempt to down-regulate flower

colour in transgenic petunia plants, and in studies on virus infection of plants, hence was

described as ‘post transcriptional gene silencing’ (PTGS) or ‘co-suppression’ (Napoli et al.

1990; Waterhouse et al. 1998). The same phenomenon has also been found in almost all

eukaryotes including mammals (Berns et al. 2004; Silva et al. 2004), insects (Kennerdell and

Carthew 2000; Somma et al. 2002), fungi (where it was termed ‘quelling’) (Cogoni and Macino

1999; Liu et al. 2002; Forrest et al. 2004) and in unicellular organisms e.g. Plasmodium and

Trypanosoma (Ngo et al. 1998; Malhotra et al 2002).

11

MiRNAs which are involved in gene silencing generate a family of ~19-25 nucleotide (nt) long

RNAs from endogenous miRNA transcripts, and these have now been found in a wide range of

organisms, from algae to plants and humans (Bartel 2004; Griffiths-Jones et al. 2008). MiRNAs

are non coding, mostly 5′ uridine containing nucleotide sequences that regulate messenger RNA

stability and translation, and hence contribute to regulating gene expression (Chendrimada et al.

2007). Some DNA viruses have also been found to encode miRNAs (Pfeffer et al. 2004), and

miRNAs in mice have also been observed to counteract viruses by targeting viral large protein

(L protein) and phosphoproteins olso called P proteins (Otsuka et al. 2007). In humans, an

endogenous miRNA, miR-32, inhibits replication of a primate retrovirus PFV-1 (primate foamy

virus type 1); thus implying miRNAs may also have direct roles in antiviral defence (Lecellier

et al. 2005).

Apart from the PTGS reported in plants, silencing of genes at the transcription level has been

studied in great detail in C. elegans (e.g. Grishok et al. 2005). This evolutionarily conserved

mechanism may have evolved as an antiviral defence mechanism, in which viral RNA is

degraded by host RNAi machinery, and by which transposon activity can also be inhibited

(Voinnet 2001; Ketting et al. 1999; Ketting and Plasterk 2000).

In C. elegans, the first miRNA identified was lin-4, which represses the translation of lin-14, a

gene involved in developmental timing during postembryonic life stages (Lee et al. 1993;

Feinbaum and Ambros 1999). Soon after a 21 nt miRNA let-7 was discovered which affects

coordination of developmental timing, and is highly conserved and expressed in late larval

stages of C. elegans (Reinhart et al. 2000; Pasquinelli et al. 2000). It is also involved in the

maintenance of heterochromatin thus indirectly controlling epigenetic gene expression (Volpe et

al. 2002; Lippman and Martienssen 2004). A genome-wide RNAi screen identified at least 19

genes involved in the miRNA pathway of C. elegans (Parry et al. 2007). As many as 54 miRNA

families are reported as conserved in four Caenorhabditis species i.e. C. elegans, C. briggsae,

C. remanei and C. brenneri (Shi et al. 2013). In plants, miRNAs have been shown to respond to

biotic stresses such as fungal and viral infections, for example 10 miRNAs were down-regulated

in loblolly pine when infected with rust fungus, while two miRNAs were greatly up-regulated

when Turnip mosaic virus infected Brassica rapa plants (Lu et al. 2007; He et al. 2008).

Recently, 40 families of miRNAs in soybean belonging to conserved miRNA families as well as

soybean specific miRNAs have shown differential expression patterns in response to soybean

cyst nematode infection (Li et al. 2012). These studies demonstrate an essential role of miRNAs

in plant defence responses in addition to gene regulation.

12

1.2.1 RNAi mechanism

The eukaryotic RNAi pathway is triggered in response to presence of double stranded (ds)RNA.

When the dsRNA is from an external source, it is referred to as an exogenous RNAi (exo-

RNAi) pathway. This pathway also includes genes encoding miRNAs which are involved in

modifying gene expression, and maintenance of the organism’s genome in which case it is the

endogenous RNAi (endo-RNAi) pathway (Correa et al. 2010). These mechanisms form the

basis of antiviral responses, epigenetic regulation, and some responses to abiotic and biotic

stresses (Ruiz et al. 1998; Vaucheret 2006; Navarro et al. 2006). DsRNA can enter the cells via

transmembrane protein transporter channels or as a result of RNA-virus infection (Saleh et al.

2006). Once inside a cell, the dsRNA is processed by the dicer complex, which includes the

ribonuclease enzyme dicer, helicases, dsRNA binding enzymes and ‘argonaute’ proteins, and

these act together to cut long dsRNAs into ~21-25 bp siRNAs. In C. elegans, an argonaute gene

rde-1, in the dicer complex appears to contribute to separating the two strands of double

stranded siRNA (Steiner et al. 2009).

The miRNA pathway is initiated when hairpin transcripts known as primary miRNAs (pri-

miRNAs), encoded in the genome, are generated by RNA polymerase II (Lee et al. 2004).

These are 5′ capped and have 3′ polyadenylated nucleotide sequences (Bracht et al. 2004). They

are processed by the enzymes drosha and its cofactor pasha into ~70 nt precursor miRNAs (pre-

miRNA) which are then exported out of the nucleus through ‘exportin’ proteins (Lee et al.

2002; Lund et al. 2004). Pre-miRNAs are taken up by the dicer complex which cleaves the ~70

nt hairpin into ~22 bp miRNAs (Grishok et al. 2001; Lee et al. 2002). The duplex miRNAs are

unwound and mature single stranded 21 bp miRNAs are then taken up by the miRNA-induced

silencing complex (miRISC), consisting of RNA binding proteins, argonautes and nucleases

(Chan et al. 2008). Similarly, the processed duplex siRNA is unwound by an RNA-Induced

Silencing Complex (RISC) in an ATP dependent manner leading to the activation of the RISC

(Nykanen et al. 2001). With the interaction of argonautes, RNA binding proteins and helicases,

RISC functions as a transport vehicle for carrying single stranded siRNA to the respective

complementary target mRNA (Tang 2005). The sequence specific base-pairing results in

mRNA cleavage at the centre of the ~21 bp siRNA which results in degradation of the mRNA

by nucleases and so prevents its translation into a polypeptide (Elbashir et al. 2001a, Elbashir et

al. 2001b). For miRNAs, the miRISC targets the complementary mRNA, usually at the 3′

untranslated region (3′ UTR), possibly because there is less chance of dislodging miRNAs by

ribosomal activity (Ruvkun et al. 1989; Gu et al. 2009). This results in translational repression

of complementary mRNAs (Figure 1.4).

13

The miRNAs together with their mRNA targets and associated argonautes co-localise in

cytoplasmic foci called ‘processing bodies’ (P-bodies) or GW bodies (Eystathioy et al. 2003).

P-bodies are involved both in mRNA degradation and mRNA storage, from where mRNA can

return to translation when required (Liu et al. 2005; Parker and Sheth 2007).

In C. elegans, plants and fungi, there is also an amplification system for small RNAs initiated

by RNA-dependent RNA polymerases (RdRps), referred to as transitive RNAi (Calo et al.

2012; Fernandez et al. 2012; Vaistij et al. 2002). In C. elegans, target mRNA is used as a

template and sense strand as the primer for the 5′ to 3′ synthesis of more dsRNAs (Alder et al.

2003). Synthesis of secondary siRNAs has also been proposed by independent or unprimed

RdRp activity resulting in a 5′ di- or tri-phosphate product, an antisense polarity and synthesis

of sequence beyond the initial trigger dsRNA (Pak and Fire 2007; Sijen et al. 2007). These

secondary siRNAs can then associate with distinct argonautes, hence repeating and amplifying

the RNAi process (Yigit et al. 2006).

Another class of small RNAs termed pi-RNAs (PIWI-interacting RNAs) is being investigated,

but currently with poorly understood mechanism and functions. Pi-RNAs are encoded by the

genome and pre-dominantly affect the gonadal tissue of nematodes, with functions affecting

epigenetic phenomena (Ashe et al. 2012). The well co-ordinated functions of small RNA

pathways in an organism require specific effectors at different steps, which can be largely

classified into functional components.

14

FIGURE OMITTED

Figure 1.4: Core components of the RNAi pathways of C. elegans. Exogenous dsRNA enters

the cells through transport proteins (1) while endo-siRNA and pre-miRNA are coded by the

organism’s genome and exported out of the nucleus (2). Both dsRNA and pre-miRNA are

processed by the Dicer complex (3) into siRNA and miRNA in the cytoplasm. After dicer

processing, these small RNAs are taken up by the RNA-Induced Silencing Complex (RISC) (5),

unwound and guided to the target mRNA resulting in target cleavage or translational repression

(6). RNAi inhibitory proteins suppress this mechanism (4) while RNAi amplification machinery

(7) generates secondary siRNAs which amplify the silencing process (8) and initiate nuclear

RNAi which leads to transcriptional silencing (10). Cell-to cell transport of silencing signals

ensures their spread throughout the organism. (Dalzell et al. 2011).

15

1.3 Components of siRNA and miRNA pathways of eukaryotes

The proteins involved in the interacting sRNA pathways of C. elegans can be divided into

groups according to their function. These include:

1. Transport proteins which allow movement of silencing signals, dsRNAs across membranes.

2. Dicer complex which unwinds and cuts longer dsRNAs into smaller RNAs required for

interacting with a target sequence.

3. RNA-induced silencing complex (RISC) which guides the siRNA and miRNA to the target

mRNA.

4. RNAi amplification machinery that amplifies gene silencing by producing secondary

siRNAs.

5. RNAi inhibitors, which include proteins that suppress gene silencing by various mechanisms

6. Nuclear RNAi is the process where silencing signals participate in silencing the nascent

mRNA of the target gene produced in the nucleus.

7. Argonautes, a group of proteins that function at different steps of gene silencing pathway

with major function of binding RNAs and cleaving target mRNA.

1.3.1 Transport proteins

These are a group of genes which encode various proteins responsible for transporting RNAs

(long dsRNA, siRNA and miRNA) from the extracellular environment across the cell

membrane and/or the endoplasmic reticulum into the cell. This group of RNA transport proteins

also includes nuclear membrane transport proteins responsible for the transport of pre-miRNA

from the nucleus to the cytoplasm.

In C. elegans, ten genes are known to encode proteins that can transport siRNA, dsRNA and/or

miRNA between cellular compartments or cells. These are sid-1, sid-2, sid-3, rsd-2, rsd-3, rsd-

6, haf-6 and the nuclear envelope transporters xpo-1, xpo-2 and xpo-3. Sid-1 is involved in the

passive uptake of dsRNA across the cell memebrane. It functions as a channel to spread the

silencing signal in the form of dsRNA (siRNA, hpRNA etc.) through cells of different tissues in

C. elegans (Jose and Hunter 2007; Shih and Hunter 2011). The 776 amino acid long

transmembrane protein SID-1 which has an extracellular amino terminus, is expressed in all

non-neuronal cells and preferentially transports long (~500 bp) dsRNAs (Winston et al. 2002;

Feinberg and Hunter 2003). However, this protein is not involved in the export of silencing

signals from cell to cell in adjacent tissues (Jose et al. 2009). In sid-1 mutants, expression of

SID-1 from cell specific promoters results in cell specific RNAi suggesting a high affinity of

dsRNAs to SID-1 (Calixto et al. 2010). SID-2 is an intestinal transmembrane protein in C.

16

elegans which aids dsRNA uptake after feeding, enabling what is termed ‘environmental’

RNAi. When expressed in environmental RNAi defective C. briggsae, sid-2 restores the

environmental RNAi response (Winston et al. 2007). SID-2 is involved in the selective uptake

of dsRNA in an ATP- and pH-dependent manner from the gut lumen (McEwan et al. 2012).

The SID-3 protein also imports dsRNA efficiently into C. elegans cells from intestinal cells and

to internal tissues, while SID-5 associates with endosomes and is involved in the efficient

spread of silencing signals out of intestinal cells independent of SID-1 (Hinas et al. 2012).

Unlike flies and vertebrates, in which pre-miRNAs are transported out of the nucleus through

exportin-5, in C. elegans it is suggested that three exportins xpo-1, xpo-2 and xpo-3 are involved

in the miRNA pathway, in which XPO-1 mediates intra-nuclear transport of pri-miRNA and

enhanced processing of pri-miRNA to pre-miRNA (Bussing et al. 2010).

Mutants of rsd-2, rsd-3 and rsd-6 in C. elegans are able to take up dsRNA from the gut into

somatic tissues, but its spread to the germline tissue is inhibited - this indicates that these genes

are important in germline RNAi spread (Tijsterman et al. 2004). A yeast hybrid assay also

shows that the products of rsd-2 require interaction with rsd-6 to function properly (Tijsterman

et al. 2004). The RSD-3 protein contains the epsin amino-terminal homology (ENTH) motif

typical of cytosolic proteins involved in vesicle trafficking (Holstein and Oliviusson 2005). The

ATP binding cassette (ABC) transporter gene haf-6 is required for efficient RNAi of genes

expressed in germline and intestinal tissues in C. elegans (Sundaram et al. 2006). All these

different effectors are involved in systemic spread of RNAi through nematode tissues and

demonstrate the involvement of several mechanisms notably active membrane uptake, passive

membrane transport and endocytosis. After entering a cell, dsRNA is taken up by the

ribonuclease III Dicer complex for processing, and this aspect is discussed in more detail in the

following section.

1.3.2 Dicer complex

The ribonuclease III dicer enzyme is the main component of the Dicer complex in eukaryotes.

There are four classes of dicer-like proteins in plants, two in fungi and insects and one in

humans and nematodes. The dicer protein(s) plays a major role in recognition and processing of

dsRNA. The dicer cleaves long dsRNA into small RNA duplexes with 3′ 2-nt overhangs,

bearing 5′ phosphate and 3′ hydroxy termini (Elbashir et al. 2001a). A 210 kDa protein is the

only DICER in C. elegans, and functions in exo-RNAi, endo-RNAi and miRNA pathways: it

processes long dsRNAs into ~22-23 bp long siRNAs in vitro with the requirement of ATP

hydrolysis, which is consistent with the in vivo studies (Ketting et al. 2001).

17

The crystal structure of the Dicer protein of Giardia intestinalis has been used to predict its

interaction with dsRNA and the results suggest that the PAZ domain binds to the 3′ end of

dsRNA and that the dicer protein acts as a ‘ruler’, starting from the PAZ domain, to process

long dsRNA into small RNAs (MacRae et al. 2006). In the nematode C. elegans, this complex

requires other effectors (drh-1, drh-2, drh-3, pir-1, rde-1 and rde-4) to process the dsRNA

trigger (Duchaine et al. 2006). By regulating gene expression, dcr-1 is proposed to control the

innate immune response of C. elegans against pathogens and possibly stress responses (Welker

et al. 2007).

In plants, it is generally accepted that there is a division of labour between the four dicer-like

proteins (DCLs), with DCL-2 involved in producing 22 bp stress-related siRNAs, DCL-3

producing 24 bp siRNAs is involved in RNA-directed DNA methylation and heterochromatin

formation, while DCL-4 is involved in post transcriptional gene silencing and produces 21 bp

trans-acting siRNAs (tasiRNAs). DCL-1 is involved in miRNA processing, whilst antiviral

responses mostly require action of DCL-2 and DCL-4 (Xie et al. 2004). These distinct sizes of

siRNAs (21 bp to 24 bp) have been documented as involved with different facets of silencing

i.e. short siRNAs guide mRNA degradation, whereas systemic silencing and DNA methylation

are responses to longer siRNAs (Hamilton et al. 2002). In Drosophila, Dicer-1 processes

miRNAs with the help of its cofactor Loquacious, while Dicer-2 processing results in siRNAs

(Saito et al. 2005). Dicer-2 also appears to have a downstream role where it interacts with RISC

with the aid of dsRNA-binding protein R2D2 (Liu et al. 2003).

In humans and drosophila, Drosha-1 RNaseIII enzyme processes pri-miRNAs into 70 nt stem

loop pre-miRNAs in the nucleus with the help of a cofactor Pasha-1, which is an RNA binding

protein required for the process (Lee et al. 2003; Filippov et al. 2000). In plants however, pri-

miRNAs, which are hairpin precursors encoded in the genome, are processed into < 100-900 nt

long pre-miRNAs and ultimately miRNA-miRNA duplex by DCL1 together with the proteins

HYPONASTIC LEAVES1 (HYL1) and SERRATE (SE), which catalyse this process in the

nucleus (Kurihara et al. 2006; Dong et al. 2008; Cuperus et al. 2011). These and siRNAs

processed by dicer then enter the RISC for further processing and eventual disruption of gene

expression.

1.3.3 RNA-induced silencing complex (RISC)

The RISC is one of the most important components of RNAi. In Drosophila one of its roles is in

retaining the dsRNA that enters the cell and maintaining an unprocessed pool of dsRNA (Shih

and Hunter 2011). The complex includes many gene products - helicases, argonautes and DNA

18

and RNA binding effectors. This ribonucleoprotein complex can load siRNAs (siRISC) or

miRNAs (miRISC, Tang 2005).

The RISC recognises the 2 nt overhang at the 3' end of the guide strand of siRNAs, binds and

unwinds it from the 5′ end into single-stranded RNA which then guides the RISC for mRNA

degradation (Chiu and Rana 2002; Kennedy et al. 2004). An additional feature of the plant

small RNA pathway is methylation of small RNA duplexes to protect them from degradation. In

plants, the 2′ hydroxyl group on the 3′ end is methylated by the protein HEN1 (HUA

ENHANCER 1) to prevent degradation with preference for 21-24 nt miRNAs and 23-24 nt

siRNA duplexes (Yang et al. 2006). Some plant viruses have evolved inhibitors of gene

silencing which inhibit HEN1 activity, resulting in degradation of siRNAs generated as a

defence against those viruses (Jamous et al. 2011). The thermodynamic properties of the two

ends of duplex siRNA and miRNA determine which strand will act as guide for RISC

(Khvorova et al. 2003; Schwarz et al. 2003). Complementarity with target mRNA at the 5′ end

is essential for translational repression while the 3′ end contributes to the establishment of an A-

form helix which is essential for RISC mediated target cleavage (Chiu and Rana 2003; Haley

and Zamore 2004). Gene regulation through miRNAs in plants results in both translational

repression and endonucleolytic cleavage of mRNAs (Brodersen et al. 2008; Huntzinger and

Izaurralde 2011).

The gene tsn-1, a component of the RISC complex, encodes six functional protein domains i.e.

four SNase (Staphylococcal nuclease) domains followed by a Tudor domain, which is

implicated in protein–protein interactions. A fifth SNase domain is present at the amino

terminus of the Tudor domain and this gene has nucleolytic activity (Caudy et al. 2003).

Another component of the ~250 kDa complex is vig-1 which in C. elegans immunoprecipitates

with tsn-1 protein and let-7 miRNA, indicating that it is also associated with the miRNA

pathway (Caudy et al. 2003). The endonucleolytic activity of the RISC requires the presence of

Mg2+

ions (Schwarz et al. 2004). In mammals, siRNAs behave as miRNAs if there is less

complementarity at the 3′ end (Doench et al. 2003).

The RISC also comprises of two effectors ain-1 and ain-2, which encode an M domain (M

domain of the protein GW182) thought to promote translational repression and initiate gene

silencing (Zekri et al. 2009). Insects like Drosophila, have one GW182 protein, while C.

elegans has two, out of which, AIN-1 interacts with miRISC and localizes to P-bodies (Ding et

al. 2005; Eulalio et al. 2007). The genes ain-1 and ain-2 appear to function in a redundant

manner in association with miRNA argonautes alg-1 and alg-2 to form distinct miRISC and

regulate expression of target mRNAs (Zhang et al. 2007).

19

1.3.4 RNAi amplification machinery

In C. elegans, an amplification machinery uses the target mRNA as a template to synthesise

more dsRNAs by employing RdRPs. The synthesis of dsRNA can continue beyond the initial

trigger sequence (Pak and Fire, 2007; Sijen et al., 2007). This phenomenon is also present in

plants and fungi, but has not been found in Drosophila or mammals: it has been termed

‘transitive RNAi’. Distinct secondary siRNAs are generated by individual RdRp events and can

be distinguished from primary siRNAs by the presence of di- or tri-phosphates at the 5′ end,

instead of mono-phosphate for primary siRNAs (Pak and Fire 2007; Sijen et al. 2007).

Seven genes are known to be involved in RNAi amplification in C. elegans. Two of these are

RdRps ego-1 and rrf-1. The gene ego-1 is important for fertility in C. elegans and is required for

efficient germline RNAi response (Smardon et al. 2000) whereas rrf-1 is required for efficient

RNAi in somatic tissues suggesting there are distinct roles for these two closely linked genes

(0.9 kb apart in tandem orientation) in C. elegans (Sijen et al. 2001).

Three C. elegans genes with the phenotypic designation “Suppressor with Morphological effect

on Genitalia” (smg-2, smg-5 and smg-6) are also known to contribute to amplification of RNAi,

however the specific mechanisms have not been elucidated. These SMG proteins are involved in

degrading defective mRNAs, those that code for toxic protein fragments and nonsense mediated

mRNA decay (also termed as mRNA surveillance, Pulak and Anderson 1993; Johns et al.

2007). Two recently characterised RNAi amplification genes of C. elegans, rde-10 and rde-11

are essential for accumulation of secondary siRNAs in endo and exo-RNAi pathways and act by

forming a complex that interacts with partially degraded target mRNAs (Zhang and Ruvkun

2012).

1.3.5 RNAi inhibitors

In contrast to proteins involved in RNAi amplification, a group of proteins known as RNAi

inhibitors appear to suppress RNAi in C. elegans: the loss of function of these increases

sensitivity to RNAi. So far 13 genes are reported to have a role in inhibiting RNAi directly or

indirectly. These are genes with enhanced RNAi phenotype (eri-1, eri-3, eri-5, eri-6/7, eri-9),

adenosine deaminase acting on RNA genes (adr-1, adr-2), XRN (mouse/S. cerevisiae)

ribonuclease related genes (xrn-1, xrn-2), RdRp containing (rrf-3), lin15-b, gfl-1 and zfp-2.

The nuclease domain of eri-1 is responsible for removing the 2 nt overhang by recognizing the

3′ end of siRNA, also known as 3′ exoribonucleolytic trimming, and is common to E. coli and 3′

processing of 5.8S ribosomal RNA in C. elegans (Li et al. 1998; Gabel and Ruvkun 2008). The

possible inhibition of RNAi by eri-1 is suggested because of loss of recognition by RISC and

20

instability of siRNAs without the overhangs, making them more susceptible to degradation

(Kennedy et al. 2004). The eri-3 and eri-5 genes are essential for stable interaction of eri-1 with

dcr-1, while eri-9 is involved in endogenously expressed siRNAs (Duchaine et al. 2006;

Pavelec et al. 2009). In C. elegans, two separate mRNAs eri-6 and eri-7 are trans-spliced to

form the RNAi inhibitor eri-6/7 which codes for a helicase that functions in both exo and endo-

RNAi pathways (Fischer et al. 2008). This suggests that in C. elegans, the same protein can

play overlapping roles in sRNA pathways and affect stability of interactions between other

effectors of the pathway.

The RdRp coding gene rrf-3 is also an inhibitor of RNAi in C. elegans since its silencing results

in increased efficiency of RNAi (Simmer et al. 2002). As a result, rrf-3 mutants of C. elegans

have been used in various RNAi experiments to study sensitivity of genes to RNAi.

Interestingly, ego-1 and rrf-1, also RdRps play exactly the opposite role, i.e. are involved in

RNAi amplification. Further functional protein analysis is needed to understand the contrasting

roles of this protein.

Two deaminating proteins are also involved in RNAi inhibition i.e. ADR-1 and ADR-2 in C.

elegans which suppress RNAi by deaminating dsRNA synthesised from transgenes and the

mutants of these genes are defective in chemotaxis (Knight and Bass 2002; Tonkin et al. 2002).

Similarly xrn-1 is required for embryogenesis, pointing to a role for XRN exoribonucleases in

developmental regulation (Newbury and Woollard 2004). The xrn-1 and xrn-2 genes are

involved in degradation of mRNA in P-bodies and mature miRNAs respectively, therefore

regulating miRNA homeostasis in C. elegans (Muhlrad et al. 1994; Chatterjee and Grobhans

2009).

The gfl-1 gene, a suppressor of RNAi, also has a reported role in eukaryotic transcription

(Heisel et al. 2010). The gene zfp-2 is a zinc-finger transcription factor which interacts with lin-

35Rb and acts as an RNAi inhibitor (Ceron et al. 2007). Mutants of some of these inhibitors

(eri-1, rrf-3, lin-15b) show increased viral resistance in C. elegans compared to wild-type

nematodes, indicating their powerful role in inhibiting virus-degrading small RNAs (Wilkins et

al. 2005; Schott et al. 2005). Characterisation of this group of genes in PPNs would help in

development of robust RNAi strategies for their control.

1.3.6 Nuclear RNAi

Post transcriptional gene silencing is not restricted to the cytoplasm of eukaryotic cells

(Montgomery et al. 1998). Silencing of transcripts in the nucleus or nuclear RNAi is also

responsible for heritable gene silencing in C. elegans (Burton et al. 2011). The secondary

siRNAs generated by RdRps are bound to nuclear RNAi pathway argonaute NRDE-3 (nuclear

21

RNAi defective-3) which normally resides in the cytoplasm, but once bound to the secondary

siRNAs, localises to the nucleus, and associates with the complementary nascent mRNA

together with NRDE-1 and NRDE-2 (Guang 2008). In the nucleus the NRDE-siRNA complex

interacts with another effector NRDE-2 to silence precursor mRNA (pre-mRNAs) and inhibit

RNA polymerase II transcription during the elongation phase of transcription (Guang et al.

2010). Two other effectors identified in the C. elegans nuclear RNAi pathway, nrde-1 and nrde-

4, are involved in chromatin association and methylation of histone 3 lysine 9 (H3K9me) which

is also detected in the progeny, hence the association of nuclear RNAi to heritable gene

silencing (Burkhart et al. 2011; Burton et al. 2011; Gu et al. 2012). The nrde-3 gene has also

been shown to play a role in RNAi amplification in C. elegans in which mutants are RNAi

defective and its over expression enhances RNAi (Zhuang et al. 2013). A model of nuclear

RNAi in C. elegans is presented in Figure 1.5.

FIGURE OMITTED

Figure 1.5: Model of nuclear RNAi in C. elegans. NRDE-3 binds to secondary siRNA in the

cytoplasm and transports it into the nucleus to interact with other nuclear RNAi proteins NRDE-

1 (1), NRDE-2 (2), NRDE-4 (4) to silence precursor mRNA (pre-mRNA) and inhibit RNA

polymerase II (Pol II) activity (Buckley 2013).

The gene cid-1 encodes a polymerase which in yeast is involved in RNAi dependent

heterochromatin formation and 3′ polyadenylation of mRNA in the nucleus (Stevenson and

Norbury 2006). However in C. elegans, this protein acts as a checkpoint, which in response to

DNA damage stalls cell division and cid-1 mutant nematodes show altered stress response

(Olsen et al. 2006). The ekl genes, also involved in nuclear RNAi, are classified as the

enhancers of the ksr-1 lethality because their knockdown causes a distinct rod-like larval

lethality in ksr-1 mutant C. elegans (Rocheleau et al. 2008).

22

Mutator genes mut-2, mut-7 and mut-16 are also involved in nuclear RNAi pathway in C.

elegans. The gene mut-2, also named as rde-3, codes for a polymerase β nucleotidyltransferase

which in C. elegans is essential for fertility, viability and siRNA accumulation for RNAi

induced by feeding (Chen et al. 2005). The mut-7 gene encodes a putative 3′-5′ exoribonuclease

essential for RNAi which interacts with rde-2 in C. elegans while mut-16 is involved in

transposon silencing and mutant worms are defective in RNAi (Vastenhouw et al. 2003; Tops et

al. 2005).

The RNA helicase rha-1, also a nuclear RNAi effector, is required for germline development:

RNAi of rha-1 in germ cells and its mutants have defective chromatin organization (Walstrom

et al. 2005). Three Maternal Effect Sterile (MES) proteins mes-2, mes-3 and mes-6 form a

complex which regulates germline development by repressing repetitive transgenes (Fong et al.

2002). This complex is also responsible for the epigenetic methylation of histone 3 lysine 27

(H3-K27) in adult germline and early embryos (Bender et al. 2004).

1.3.7 Argonautes

Argonautes are a large group of genes encoding proteins that have the PAZ

(Piwi/Argonaute/Zwille) and PIWI domains. They are involved in a range of small RNA

pathways and other processes including chromatin modification and stem cell fate determination

(Carmell et al. 2002). The PAZ domain binds RNA by recognizing the 2 nt 3′ overhang of

siRNAs in a sequence-independent manner (Song et al. 2004). Removal of one or both of these

overhangs reduces the binding affinity by 85-fold and >5000-fold respectively (Ma et al. 2004).

The PIWI domain of argonautes is ribonucleolytic and acts as the catalytic slicer site involved in

target slicing. The crystal structure indicates that it is an RNase H domain (Song et al. 2004;

Parker and Barford 2006).

After studying the crystal structure a model proposed for a full length argonaute of the

archaebacterium P. furiosus, suggests 3′ end of siRNA binds to the PAZ cleft and the target

mRNA is oriented so that the PIWI RNase H site cleaves it between the 11th and 12

th nucleotide

counting from the 3′ end of the guide siRNA (Figure1.6). Studies of the PIWI domain reveal

that 5′ phosphate-containing RNA is bound more tightly to the domain compared to a non

phosphorylated RNA, and this 5′ end initiates base-pairing with the target mRNA (Martinez et

al. 2002; Haley and Zamore 2004; Patel et al. 2006).

23

FIGURE OMITTED

Figure 1.6: Proposed mechanism of the slicer activity of P. furiosus argonaute on siRNAs. The

3′ end of the siRNA binds to the PAZ cleft and is oriented in a position which allows the PIWI

RNaseH to cleave the siRNA between 11th and 12

th nucleotide (Song et al. 2004)

The genome of C. elegans codes for 28 argonaute proteins whilst in Drosophila, Arabidopsis

and humans the argonaute functions are divided amongst 5, 10 and 8 orthologs respectively.

Some important argonautes in different eukaryotes and their functions are presented in Table

1.2.

Table 1.2: Argonaute orthologs, their respective functions and mutant phenotypes in model

organisms.

Argonaute Proposed function Mutant phenotype Reference

Nematodes (C. elegans)

RDE-1 Processing of duplex primary

siRNAs i.e. removal of

passenger strand from guide

strand

RNAi defective Tabara et al. 1999

Steiner et al. 2009

ALG-1 and ALG-2 Interaction with miRNAs with

functional homology to rde-1

Inhibited miRNA

processing

Grishok et al. 2001

Tops et al. 2006

ERGO-1 Homologous in function to rde-

1 in the endo-RNAi pathway

RNAi sensitive Yigit et al. 2006

CSR-1 Involved in chromosome

alignment and segregation.

Also associated with secondary

siRNAs

Sterile Yigit et al. 2006

Aoki et al. 2007

Claycomb et al. 2009

PRG-1 Required for the presence of

21U-RNA which affect

spermatogenesis transcripts

Sterile Yigit et al. 2006

Wang and Reinke

2008

SAGO-1/WAGO-8

and SAGO-2/

WAGO-6

Interaction with secondary

siRNAs

Reduced RNAi activity Yigit et al. 2006

PPW-1 Interaction with secondary

siRNAs and involved in

germline RNAi

Resistant to RNAi of

germline expressed

genes

Yigit et al. 2006

Tijsterman et al.

2002

PPW-2/WAGO-3 Exo and endo-RNAi including

transposon silencing

Resistant to germline

RNAi

Vastenhouw et al.

2003

Gu et al. 2009

F58G1.1/WAGO-4 Interaction with secondary

siRNAs and involved in

germline RNAi

Resistant to germline

RNAi

Yigit et al. 2006

Gu et al. 2009

24

C16C10.3/HRDE-1 Endogenous RNAi leading to

nuclear gene silencing in germ

cells

Heritable RNAi

deficient

Buckley et al. 2013

NRDE-3/WAGO-12 Nuclear RNAi Nuclear RNAi defective Guang et al. 2008

Plants (A. thaliana)

AGO1 Processing of pre-miRNAs and

guiding the processed miRNA

to target mRNA

Upregulation of miRNA

target gene transcripts

Baumberger and

Baulcombe 2005

Vaucheret et al. 2004

AGO2 Essential for antiviral defence Decreased antiviral

defence response

Carbonell et al. 2012

AGO3 Unknown No obvious phenotype Lobbes et al. 2006

AGO4 Involved with chromatin

silencing

Defective in DNA

methylation and

accumulation of 25 bp

siRNAs

Zilberman et al.

2003

AGO5 Binds to 21-nt miRNa and 21

and 24 nt viral siRNAs. -

Takeda et al. 2008

AGO6 Involved with RNA-directed

DNA methylation and

transcriptional gene silencing in

meristematic tissues

Suppressed RNA

silencing

Zheng et al, 2007

Eun et al. 2011

AGO7 Interaction with miRNAs to

stabilise target interaction and

tasiRNA biogenesis

Decreased tasiRNA

biogenesis. Abnormal

development.


AGO9 Involved with RNA-directed

DNA methylation

Defective gametogenesis Olmedo-Monfil et al.

2010

AGO10/PINHEAD/Z

WILLE

Interaction with miRNAs to

stabilise target interaction,

maintenance of meristematic

cells in shoot apex

Defective meristem

formation

Moussian et al. 1998


Tucker et al. 2008

Insects (D. melanogaster)

AGO-1 Involved in miRNA pathway Up-regulation of

miRNA targets and

decreased concentration

of miRNAs

Forstemann et al.

2007

Okamura et al. 2004

AGO-2 Involved in siRNA-directed

RNAi

RNAi defective embryos Forstemann et al.

2007

Okamura et al. 2004

Fungi (N. crassa)

QDE-2 siRNA processing. Involved in

processing miRNAs from pre-

miRNAs

RNAi defective Maiti et al. 2007

In a mutant screen, C. elegans argonaute functions were explored with the following findings

(Yigit et al. 2006)

1. Argonautes are involved in nematode fertility and chromosome segregation.

2. Different argonautes act at different steps in sRNA pathways.

3. Exo-RNAi and endo-RNAi pathways converge, probably on SAGO proteins, and

compete for the same resourcess.

25

Ten argonaute proteins have been identified in Arabidopsis. Unlike animals, after being

exported out of the nucleus, it is proposed that pre-miRNAs are recruited by AGO1 instead of

RISC, which carries out the slicing activity and are guided to the imperfect complementary

target mRNAs (Baumberger and Baulcombe 2005). AGO1 has a preference for small RNAs

with a 5′ terminal uridine which is a characteristic of most miRNAs. The AGO1 mutants in

Arabidopsis show up-regulation of mRNAs which are known targets of miRNAs (Vaucheret et

al. 2004; Mi et al. 2008). The 5′ terminal nucleotide of the small RNA plays a significant role in

determining which Argonaute protein they will interact with. For example, AGO2 and AGO4

preferentially interact with RNAs with a 5′ terminal adenosine while AGO5 binds small RNAs

starting with a cytosine (Mi et al. 2008). Plant viruses and pathogenic fungi have evolved

methods to hijack AGO1 of plants (e.g. Arabidopsis) to disarm the plant’s immune response and

achieve successful infection (Azevedo et al. 2010; Weiberg et al. 2013). Although argonautes

are known for their ‘slicer’ activity, they also play major roles in the miRNA pathway where

target slicing does not occur. The fact that in C. elegans, a single Dicer processes small RNAs

which are then taken up by different argonautes, while in plants Dicer processing properties

decide the specific argonaute association, indicates the diversity of the roles these proteins play

in the sRNA pathways.

1.4 Applications of RNAi

Nominated as ‘Breakthrough of the Year 2002’ by the journal Science, RNAi has since been

studied intensively, because it presents new approaches to counteract chronic diseases in

medical science, and has many potential contributions in crop improvement. Medical

applications are commonly known as ‘RNAi therapeutics’. RNAi has been shown to repress

hepatitis B viral RNA by introducing hepatocyte targeted siRNAs. Clinical trials have been

initiated for this study (Wooddell et al. 2013). RNAi has also been used to limit replication of

HIV-1 in human cells (Jacque et al. 2002; Lee et al. 2002), and antisense oligonucleotide drugs

which share the same basic principle of targeting complementary mRNA are being developed

and clinical trials are ongoing to control some forms of cancer (Gleave and Monia 2005;

Lightfoot and Hall 2012). An antisense oligo drug ‘Fomivirsen’ has now been approved by the

FDA for the treatment of Cytomegalovirus (CMV) retinitis (Grillone and Lanz 2001).

Nanoparticle encapsulated siRNAs have demonstrated to treat 100% of rehses monkeys infected

by Makona strain of Ebola indicating the potential this technology has against treating life

threatening disease outbreaks (Thi et al. 2015).

Applications of RNAi in agriculture are being explored, for example to develop new methods to

control economically damaging pests such as nematodes and aphids. In host mediated RNAi,

gene silencing results when a parasite takes up dsRNA from the host on feeding. Experimental

26

results suggest that RNAi can be effective against both chewing and sucking insect pests (Mao

et al. 2007; Thakur et al. 2014), viruses (Tenllado et al. 2004; Bonfim et al. 2007), parasitic

weeds (de Framond et al. 2007; Aly et al. 2009) and some fungi (Nowara et al. 2010; Tinoco et

al. 2010). Topical application of dsRNA has been demonstrated to have potential to control

mosquitoes (Pridgeon et al. 2008). DsRNA foliar application against Colorado potato beetle has

also shown promising results in greenhouse conditions (San Miguel and Scott 2015). Antisense

oligonucleotides are also being tested as potential pesticides and methods of delivery are being

developed to efficiently target pests (Tortora et al. 2011).

1.4.1 Development of in vitro RNAi in nematodes

After the initial experiments in which the phenomenon of RNAi was elucidated in C. elegans,

many researchers have explored the potential of this strategy for studying functional genomics.

Gene silencing in C. elegans was first achieved by injecting dsRNA into the nematodes (Fire et

al. 1998). While exploring several methods to deliver dsRNA into these nematodes, it was

established that apart from injecting dsRNAs, environmental RNAi i.e. soaking or feeding the

nematodes with dsRNA also results in specific gene silencing (Tabara et al. 1998; Maeda et al.

2001). It was then found that feeding C. elegans with bacteria expressing dsRNA

complementary to endogenous target genes also resulted in efficient RNAi, which worked very

well for epithelial cells (Timmons et al. 2001). However, the efficiency of RNAi treatments

varied from species to species (Winston et al. 2007). Nevertheless, since the discovery of RNAi,

it has been used in studies to determine gene functions by down-regulating the expression of

about 86% of the predicted genes in C. elegans, including many genes involved in the RNAi

pathway (Fraser et al. 2000; Gonczy et al. 2000; Kamath et al. 2003; Kim et al. 2005).

Significantly, in many cases the information generated from these studies can be mapped

directly across to PPNs using a comparative genomics approach.

However, some of the methods that work for C. elegans in RNAi experiments do not work the

same way for PPNs. Microinjection of dsRNA is much more difficult, and the gene mutants

available for C. elegans are not available for PPNs, and so functional genomics studies require

alternative methods to effectively knock out genes for functional studies. Nevertheless, the

wealth of functional data generated for C. elegans can in many cases be mapped across to PPNs,

and this information has provided a major new resource to study PPNs.

In considering some other aspects of RNAi in C. elegans, which provide leads for study of

PPNs, systemic transmission of a silencing signal i.e. the ability for silencing to spread through

the organism except neuronal tissue does occur in C. elegans (Fire et al. 1998). In addition, the

inheritance of RNAi induced phenotypes lasts for up to two generations, while transcriptional

27

silencing effects can persist for many generations (Grishok et al. 2000; Vastenhouw et al.

2006). Multiple gene RNAi has also been demonstrated (Geldhof et al. 2006; Tischler et al.

2006; Gouda et al. 2010), that is, simultaneous down regulation of more than one target gene. In

contrast, specifically for animal parasitic nematodes, the efficiency of RNAi also depends on the

stability and level of transcript as well as the site of gene expression. The success rate for RNAi

is greater for genes expressed in the intestine, excretory cells and amphids than in the nervous

system (Samarasinghe et al. 2011). There are now a series of successful reports of gene

knockdown using RNAi in PPNs, but insensitivity of some genes to RNAi of these pests has

also been reported (Wheeler et al. 2012).

1.4.2 In vitro RNAi of PPN genes

PPNs are relatively difficult organisms to maintain and culture, and uptake of dsRNA was

difficult to achieve, hindering study of RNAi effects initially. A significant advance came with

the demonstration by Urwin et al. (2002) that PPNs could be induced to take up exogenous

dsRNA in solution with the addition of the neurostimulant octopamine, which induces

pharyngeal pumping in J2 worms so that they take up the external solution. This method known

as ‘soaking’ works reasonably well. Other components of soaking solutions can be added to

increase the efficiency of uptake of dsRNA in feeding experiments; these are spermidine and

gelatine. Other neurostimulants such as serotonin and resorcinol have also been used to

stimulate swallowing. Serotonin appears to cause pharyngeal contractions, and resorcinol

induces pharyngeal pumping, resulting in increased uptake of fluorescein isothiocyanate (FITC)

when used as a marker for ingestion of the external solution. However, prolonged exposure to

resorcinol is deleterious to M. incognita for incubation times longer than 4 hours (Rosso et al.

2005), and root lesion nematodes clearly do not tolerate it (Tan et al. 2013). There have been a

series of studies on in vitro feeding of PPNs targeting different genes, either individually or in

combination. These are summarised in Table 1.3.

Table 1.3: A summary of RNAi studies on PPN genes via in vitro RNAi and effects on

nematode infectivity and development.

Gene description Gene name or

Symbol

Nematode

species

Phenotype/Effect Reference

C-type lectin hgctl H. glycines Reduced transcript level.

41% less infection.

Urwin et al. 2002

Major Sperm Protein msp H. glycines Reduced transcript level

Cysteine proteinase Gpcp-I G. pallida A shift in sexual fate

(21% less females)

Hgcp-I H. glycines A shift in sexual fate

(25% less females)

Esophageal gland

proteins

Mi-crt

M. incognita

65% reduction in

transcript level

Rosso et al. 2005

Mi-pg-1 30% reduction in

28

transcript level

Cuticle and egg

development

Chitin synthase M. artiellia Defective hatching Fanelli et al.,2005

Extracellular matrix

proteins

Dual oxidase M. incognita 70% reduced egg mass

production

Bakhetia et al.

2005

Multiple functions Aminopeptidase H. glycines 61% reduced infection

on soybean roots

Lilley et al. 2005

Cell wall degrading

enzyme

Β-1,4,

endoglucanase

G. rostochiensis Reduced infection Chen et al. 2005

Amphid secreted

protein

ams-1 Reduced root invasion

Parasitism 16D10 M. incognita 74-81% reduced

infection

Huang et al. 2006

Ribosomal Protein Hs-rps-23 H. glycines Lethal Alkharouf et al.

2007

Larval molting Cathepsin L

cysteine

proteinase

M. incognita 60% less infection Shingles et al.

2007

Pharyngeal gland cell

proteins

hg-eng-1, hg-

syv46

H. glycines Reduced plant infection

establishment

Bakhetia et al.

2007

hg-gp, hg-cm,

hg-pel

Increased male to female

ratio

FMRFamide-like

Neuropeptide

Gp-flp-1, Gp-

flp-6, Gp-flp-12,

Gp-flp-14 and

Gp-flp-18

G. pallida Defective locomotion,

motor dysfunction and

increased neuronal

RNAi

Kimber et al. 2007

Esophageal gland

protein (Pectate

lyase)

Mi-gsts-1 M. incognita Decreased egg mass

production and 90%

reduction in transcript

level

Dubreuil et al.

2007

Esophageal gland

protein

Hspel2 H. schachtii 50% less infection and

decreased transcript

level

Vanholme et al.

2007

Esophageal gland

protein

Hg-pel-1

H. glycines

203-fold decrease in

transcript level

Sukno et al. 2007

Hg-4E02 51-fold decrease in

transcript level

Avirulence effector Cg-1 M. javanica Increased virulence on

tomato with Mi-1

Gleason et al.

2008

Dorsal pharyngeal

gland proteins

Dg13

H. glycines

High male to female

ratio.

Bakhetia et al.

2008

Dg14 High male to female

ratio.

Dg21 High nematode

establishment and male

to female ratio.

Esophageal gland

protein

Cellulose

binding protein

M. javanica 53-58% reduction in egg

mass production

Adam et al. 2008

Muscle (motor

proteins)

Bx-myo-3 and

Bx-tmy-1 B. xylophilus

Defective locomotion Park et al. 2008

Heat shock protein Bx- hsp-1 Lethality due to heat

sensitivity

Esophageal gland

protein

(endoxylanase)

Rsxyl R. similus 53-66% reduction in

plant infection

Haegeman et al.

2009

FMRFamide-like

Neuropeptide

Gp-flp-12 G. pallida Decreased transcript

level and inhibition of

migratory ability

Dalzell et al.

2010a Mi-flp-18 M. incognita

29

Microprocessors of

miRNAs

drsh-1 and

pash-1

M. incognita Lethal and abnormal

embryo

Dalzell et al.

2010b

Cell death

programming gene

Mi-ced-9 M. incognita 40% reduction in gall

formation

Gaeta et al. 2011

Esophageal gland

protein (Calreticulin)

Micrt-1 M. incognita Reduction in transcript

and plant infection

Arguel et al. 2012

Zinc finger protein Mi-Pos-1 M. incognita 85% reduced transcript

40% reduced hatching

Matsunaga et al.

2012

FMRFamide-like

Neuropeptide

Gp-flp-32 and

Gp-flp-32R

G. pallida 55% and 75% reduction

in transcript respectively

Atkinson et al.

2013

Muscle troponin C Mg-Pat-10 M. graminicola 91.8% inhibition in

mobility

Nsengimana et al.

2013

Muscle myofilament

calponin

Mg-Unc-87 M. graminicola 87.9% inhibition in

mobility

Muscle troponin C Pzpat-10 P. zeae 3.6-fold reduction in

transcript level

Tan et al. 2013

Muscle myofilament

calponin

Ptunc-87 P. thornei 29.9-fold reduction in

transcript level.

Pharyngeal gland cell

proteins

(endoglucanase)

Pv-eng-1 P. vulnus 88-98% reduced

transcript level. 54%

reduction in

reproduction rate.

Fanelli et al. 2014

In most cases in vitro feeding studies of dsRNA for single genes have shown promising results.

On the other hand, when dsRNAs of two or more different target genes are delivered together to

nematodes, it rather surprisingly results in less target suppression compared to when individual

dsRNAs were delivered (Bakhetia et al. 2008). Major focus of previous work has focussed on

knockdown of genes thought to be essential for parasitism and to understand effector

function(s). However, there has not been much work undertaken on down-regulation of genes

involved in the RNAi pathway: the one only study reported involved RNAi of drsh-1 and pash-

1 of M. incognita (Dalzell et al. 2010b).

1.4.3 Application of virus-induced gene silencing to PPN research

In addition to soaking in dsRNA and engineering plants to deliver gene silencing triggers

(Section 1.4.5), virus-induced gene silencing (VIGS) provides an additional approach to deliver

dsRNA triggers to nematodes in a transient manner. In VIGS, a virus is used as a vector to carry

cDNA of the gene of interest. While replicating its RNA, the virus replicates the target RNA

and so generates dsRNA to the cDNA sequence. Virus infection of cells therefore delivers

dsRNA of target genes to those cells. This dsRNA is processed by the plant antiviral machinery

to generate siRNAs, and this could result in targeting mRNA of the feeding nematode. VIGS is

one way to undertake a quick analysis of functions of nematode genes when taken up from a

plant. It has also been used to silence plant genes to study their functions without going through

lengthy and time consuming process of plant transformation (Burch-Smith et al. 2006).

30

Tobacco rattle virus (TRV) carrying M. incognita troponin C and calreticulin genes when

inoculated to tobacco plants induced dsRNA and siRNA triggers. RKNs infection of those

plants showed up to 90% reduction in transcript levels in eggs and 75% in the progeny. J2

nematodes of the progeny showed 63.5% and 84% fewer galls respectively when infected on

tomato plants (Dubreuil et al. 2009). A similar strategy was adopted for the M. javanica

NULG1a gene which codes for a secretory effector protein expressed in esophageal glands. It

resulted in a reduction of nematode infection by up to 88.8% on TRV infected tomato plants

(Lin et al. 2012).

Similarly, VIGS of the M. incognita mitochondrial ATP synthase b subunit gene (MIASB) in

tomato seedlings resulted in 64% fewer galls than controls on infection. These observations

suggest that the dsRNA triggers induced by VIGS can be taken up by nematodes, and

transported from the gut to target the gene for knockdown (Huang et al. 2014). VIGS can be

used to study the effects of gene silencing on PPNs or in systems they interact with, but VIGS is

not appropriate for developing resistance on a commercial basis.

1.4.4 The potential for PPN control using RNAi

Classical breeding for pest resistance is time consuming and requires large scale screening of

gene pools to find resistant genotypes. Achieving tolerance against plant pests/parasites through

transgenics is an attractive alternative approach because of its specificity and ability to work

with genes of multiple organisms. Inherent gene silencing mechanisms in plants allow an

appropriate transgene to produce dsRNA and convert it to siRNAs. This approach has been

explored to study the potential for host induced gene silencing (HIGS) and derived resistance to

control plant pests. This makes HIGS an exciting prospect for developing resistance to PPNs,

and research results support this concept as a viable alternative to provide host resistance for

crop protection. In contrast, transcriptional silencing of the dsRNA construct has also been

reported at variable levels both within and between T3 transgenic events in transgenic

Arabidopsis against nematodes (Kyndt et al. 2013).

HIGS is being studied from a number of angles, for example to evaluate long term effects of

gene knockdown and to assess the effectiveness of different constructs and target genes. Many

of such HIGS studies have made use of the model plant Arabidopsis, because it is a host for cyst

and root-knot nematodes (Sijmons et al. 1991; Wyss and Grundler 1992). However, in addition

to studies using plants like Arabidopsis and tobacco, HIGS has also been achieved for crop

plants like soybean, tomato and potato. A summary of the results available are provided in

Table 1.4.

31

Table 1.4: A summary of experimental studies using HIGS to control PPNs.

Plant Gene targeted Nematode

species

Effect of host

delivered knockdown

Reference

Arabidopsis

Secreted peptide

(16D10)

M. incognita 63% reduction in

infection and 69-83%

reduction in egg

production

Huang et al. 2006

Secreted peptide

(16D10)

M. javanica 63% reduction in


reduction in egg

production

Secreted peptide

(16D10)

M. arenaria 63% reduction in


reduction in egg

production

Secreted peptide

(16D10)

M. hapla 63% reduction in


reduction in egg

production

Arabidopsis

Secreted peptide

(16D10L)

M. chitwoodi 68-74% reduction in

egg mass production

Dinh et al. 2014

Nematode effector

(NULG1a)

M. javanica 88% reduction in

infection

Lin et al. 2012

Ubiquitin-like

protein (4G06)

H. schachtii

23-64% reduction in

developed females

Sindhu et al. 2009

Cellulose binding

protein (3B05)

12-47% reduced

infection

SKP1-like protein

(8H07)

>50% reduced

infection

Zinc finger protein

(10A06)

42% reduced infection

Nematode secreted

peptide, Hssyv46

36% reduced cyst

formation

Patel et al. 2008

Nematode secreted

peptide, Hs5D08

20% reduced cyst

formation

Nematode secreted

peptide, Hs4E02

20% reduced cyst

formation

Nematode secreted

peptide, Hs4F01

55% reduced cyst

formation

Parasitism effector

(30C02)

92% reduced cyst

formation

Hamamouch et al.

2012

Parasitism gene

(Mi8D05)

M. incognita Up to 90% reduction in

infection

Xue et al. 2013

Tobacco

SNF chromatin

remodelling

complex component

(snfc-5) M. incognita

>90% reduction in

infection

Yadav et al. 2006

Pre-mRNA splicing

factor (prp-21)

>90% reduced

infection

Putative

transcription factor

MjTis11

M. javanica Decreased transcript

level in feeding

nematodes

Fairbairn et al.

2007

FMRFamide-like

gene (flp-14) M. incognita

Up to 86% reduction in

reproduction rate

Papolu et al. 2013 FMRFamide-like

gene (flp-18)

Up to 82% reduction in

reproduction rate

Serine protease gene

(ser-1)

30% reduced egg

production

de Souza Junior et

al. 2013

32

Cysteine protease

(cpl-1)

42% reduced egg

production

Soybean

Major sperm protein

H. glycines

68% reduction in cyst

formation

Steeves et al. 2006

Ribosomal protein

3a(rps-3a)

87% reduced cyst

formation

Klink et al. 2009

Ribosomal protein 4

(rps-4)

81% reduced cyst

formation

Spliceosomal SR

protein (spk-1)

88% reduced cyst

formation

Synaptobrevin (snb-

1)

93% reduced cyst

formation

Beta subunit of the

COPI complex

(Y25)

81% reduced cyst

formation

Li et al. 2010

Pre-mRNA splicing

factor (prp-17)

79% reduction in

infection

Uncharacterised

protein (cpn-1)

95% reduction in

infection

L-Lactate

dehydrogenase

M. incognita

57% reduced infection,

77% reduction in

nematode diameter

Ibrahim et al. 2011

Mitochondrial

stress-70 protein

(2% reduction in

infection,8% reduced

nematode diameter

ATP synthase beta-

chain mitochondrial

precursor


62% reduced nematode

diameter

Tyrosine

phosphatise


82% reduced nematode

diameter

Tomato

Troponin C (snc)

M. incognita

59% reduced hatching

from eggs Dubreuil et al.

2009

Calreticulin (crt) No effect after

silencing. Reduced

infection by the

progeny

Dual oxidase (duox) 61% reduced infection,

52% reduction in

saccate nematodes Charlton et al. 2010

Signal peptidase

complex 3 (spc3)

52% reduction in

infection, 63%

reduction in saccate

nematodes

Subunit of 19S

regulatory complex

(rpn-7)

Up to 66.5% reduction

in infection Niu et al. 2012

Subunit of 19S

regulatory complex

(rpn-7)

50.8% reduction in egg

production

Potato Secreted peptide

(16D10L)

M. chitwoodi 44-56% reduction in

egg mass production

Dinh et al. 2014

Grape (Hairy root) 16D10 M. incognita Significant reduction

in egg production

Yang et al. 2013

33

In addition to RNAi-based transgenic resistance, alternative transgenic strategies have also been

studied. One example of these is using nematode gut protease inhibitors, for example the rice

oryzacystatin-I gene, which conferred resistance when it was transformed into tomato hairy

roots: it also strongly reduced growth and development of G. pallida (Urwin et al. 1995). A

modified rice cystatin gene (oryzacystatin-IΔD86) expressed in A. thaliana reduced growth and

fecundity of both CNs and RKNs because of inhibition of cysteine proteinase activity in the

nematode intestine (Urwin et al. 1997a). Transgenic potato lines expressing a cysteine

proteinase inhibitor from sunflower showed 8-60% resistance against G. rostochiensis and G.

pallida (Urwin et al. 2003). Although extensive research is required before commercial

implementation (Fosu-Nyarko and Jones 2015), all these transgenic resistance studies indicate

great potential of this technology for crop protection against pests, to overcome problems like

the limited genetic base of natural resistance, use of harmful chemicals and hurdles of large

scale pest control.

1.5 Aims and objectives of this project

The overall aim of this project was to extend understanding and efficiency of the HIGS

approach to control PPNs. One promising new area of study is the down-regulation of genes

encoding proteins involved in the RNAi pathway, since these might well result in lethality or

loss of function and/or ability of the nematode to parasitise its host. Arabidopsis is a host for

RKNs, and as a model plant which can be transformed readily, it was chosen as a host plant for

this study. Thus the aim of this project was to evaluate the effects of RNAi of genes involved in

the gene silencing pathway itself on the survival and parasitic ability of RKNs. Since genes of

the RNAi pathway are part of a conserved biological mechanism, the effectors of small RNA

pathways in one species could be effective targets that might be used to control a whole genus

of parasites rather than a single species.

The specific objectives of the project were:

1. To apply comparative bioinformatics and molecular tools, and information available for

C. elegans and animal parasitic nematodes, to genes involved in the RNAi pathway of

RKNs, combined with using ESTs and genomic data available for M. hapla and M.

incognita.

2. To study the effect of down-regulating expression of the identified RNAi

genes/effectors via in vitro feeding of dsRNA on the survival and/or parasitism of M.

incognita.

3. To investigate the effect of using dsRNA corresponding to different parts of an RNAi

pathway gene as a trigger for down-regulation of expression and assess the effects on

34

nematode parasitism and reproduction and relative expression of other RNAi pathway

components.

4. To investigate the possibility of controlling M. incognita via host-induced

siRNA/dsRNA of RNAi effectors.

35

The following results Chapters of this thesis are presented in the form of draft

manuscripts with the following titles, which may be modified later for publication.

Chapter 2: IQBAL, S., FOSU-NYARKO, J. & JONES M. G. K. Genome level identification

and comparison of effectors of the RNAi pathway of the parasitic nematodes Meloidogyne

hapla, Meloidogyne incognita, Ascaris suum and Brugia malayi.

Chapter 3: IQBAL, S., FOSU-NYARKO, J. & JONES M. G. K. Identification of target genes

from among sRNA pathway effectors of M. incognita for nematode control via in vitro RNAi

Chapter 4: IQBAL, S., FOSU-NYARKO, J. & JONES M. G. K. Host-induced gene silencing

of RNAi effectors confers resistance against Meloidogyne incognita and affects development.

Chapter 5: IQBAL, S., FOSU-NYARKO, J. & JONES M. G. K. The effects of RNAi

treaments with different regions of the Dicer-like gene on the viability, parasitism and

reproduction of M. incognita.

With the above format, it is the intention that ‘Introduction’ sections will be modified

for individual publication.

36

Chapter 2

Genome level identification and comparison of

effectors of RNAi pathway of the parasitic nematodes

Meloidogyne hapla, Meloidogyne incognita, Ascaris

suum and Brugia malayi.

37

Genome level identification and comparison of effectors of the RNAi

pathway of the parasitic nematodes Meloidogyne hapla, Meloidogyne

incognita, Ascaris suum and Brugia malayi.

Sadia Iqbal, John Fosu-Nyarko and Michael G.K. Jones

Plant Biotechnology Research Group, School of Veterinary and Life Sciences, WA State

Agricultural Biotechnology Centre, Murdoch University, Perth, WA 6150, Australia

2.1 Abstract

Since the discovery of RNAi as an endogenous mechanism of gene regulation in a range of

eukaryotes from unicellular organisms to mammals, it has become important to study the

underlying mechanism for this phenomenon. Because RNAi is one of the most widely used

methods for functional genomics studies, detailed information is required about the effectors

that modulate its effectiveness. The aim of this study was to identify genomic and protein

differences between the model nematode C. elegans and four parasitic nematodes i.e. Ascaris

suum, Brugia malayi, Meloidogyne hapla and Meloidogyne incognita. A further aim was to

identify the specific genomic sequence IDs coding for the genes involved in the small RNA

pathways. The results indicate interesting differences in the effectors: some are absent in

genomic sequences of parasitic nematodes, especially genes involved in the amplification and

inhibition of RNAi. There were structural differences in protein domains of effectors of the free

living and parasitic nematodes. The absence of between 38-48% of the studied effectors in the

parasitic nematodes points towards the need for detailed study of these pathways. RNAi

experiments and proteomic studies that generate information about the small RNA processes

involved in the growth and development of these parasites will help map out the RNAi pathway,

explore the viral response specific to parasitic lifestyle if any, and possibly lead to development

of more robust control strategies based on specific knowledge of RNAi for these pests.

Keywords: RNAi, miRNA pathway, genome sequence, parasitic nematodes, C. elegans.

2.2 Introduction

RNAi or post transcriptional gene silencing (PTGS) includes various small RNA pathways i.e.

exo siRNA, endo siRNA and miRNA pathways. Apart from being important in development

and regulation of gene expression, endo siRNA and miRNA pathways are responsible for

silencing viral messages and transposable elements (Ketting et al. 1999). Since the detailed

discovery of RNAi mechanisms in C. elegans, it has become a model organism to study

functional roles of various genes. Extensive studies have been conducted to understand various

38

pathways and functions using RNAi to silence different genes in C. elegans (Shelton et al.

1999; Gonczy et al. 2000; Dudley et al. 2002; Gouda et al. 2010). It was only when Urwin et al.

(2002) showed that RNAi could be induced by soaking PPNs in specific solutions containing

dsRNA that further studies using this approach demonstrated a reduction in viability,

reproduction and ultimately parasitism, depending on the genes chosen for down-regulation

(Urwin et al. 2002; Bakhetia et al. 2005; Tan et al. 2013).

In recent years, an enormous amount of data has been generated using advanced genome and

transcriptome sequencing techniques making it possible to use bioinformatics tools to study in

detail the genetics of various organisms. Belonging to the same phylum, C. elegans and

parasitic nematodes have striking anatomical similarities. However, the parasitic behaviour of

these nematodes means some genes are specific to the parasitic life style and need to be further

explored. Between 35-70% of the C. elegans genes have been reported as having homologues in

28 parasitic nematode species making it a perfect candidate to use as a model in studying

various cellular functions and pathways for which some studies have been conducted before

(Parkinson et al. 2004; Rosso et al. 2009; Dalzell et al. 2011).

The overall aim of this study was to identify genes involved in the RNAi pathway of parasitic

nematodes, and to compare and contrast the gene structures involved in these pathways between

C. elegans and parasitic nematodes and also between animal and plant parasitic nematodes.

Available genome sequences of A. suum, B. malayi, M. hapla and M.incognita were compared

to well characterized C. elegans genes involved in the RNAi pathway. This analysis led to the

identification of RNAi pathway genes in parasitic nematodes. Differences in the proteins coded

by specific genes between the different nematode genera were also explored to provide a better

understanding of the function of these genes.

2.3 Materials and methods

2.3.1 Identification of effectors of RNAi of C. elegans

Sequences of the effectors of the RNAi pathway of C. elegans were used as a primary source to

identify orthologues in the parasitic nematodes, because these have been more highly

characterised than those of any other species. Genes encoding 87 effectors directly involved in

siRNA and miRNA processes of C. elegans were identified from the literature (Rosso et al.

2009; Dalzell et al. 2011; Kikuchi et al. 2011). The nucleotide sequences of these genes were

retrieved from Wormbase version WS241 (http://www.wormbase.org) and the National Centre

of Biotechnology Information (NCBI, http://www.ncbi.nlm.nih.gov). The effectors were

grouped together, based on their functional roles in the RNAi pathway, into: those involved in

transport of silencing triggers (e.g. dsRNA, siRNA and miRNA), the Dicer and RISC

39

complexes, amplifiers of silencing signals, RNAi inhibitors, nuclear RNAi effectors and

argonautes.

2.3.2 Identification of genomic contigs of parasitic nematodes mapped to

RNAi effectors of C. elegans.

Coding sequences (mRNA) of C. elegans effectors were used to query the NCBI dataset

containing ESTs and whole genomic sequence/contigs databases of M. incognita, M. hapla, A.

suum and B. malayi with an expected value (e-value) cut-off of 10E-5. In cases where there

were no matching genomic contigs or the e-values were not significant for the C. elegans query,

sequences of effectors of parasitic nematodes were used for further confirmation in tblastx

searches: blastn and blastx were also used where required for further analysis and these are

indicated in the results. The resulting mapped contigs were retrieved with e-values, and bit

scores as an XML file and converted to csv for further analysis. To select contigs that

represented specific effectors, all matching contigs to effectors were aligned and the best match

coding for the protein domains similar to the query was used for further analysis.

2.3.3 In silico functional analysis of putative effectors of the parasitic

nematodes

Genomic contigs of any of the parasitic nematodes identified to contain RNAi effector

sequences were further analysed using a suite of bioinformatics tools. The aim was to determine

the presence of functional domains and conserved motifs that characterise specific effectors and

to determine their homology, structural and functional conservation among parasitic nematode

groups and that of C. elegans. Functional protein domains of these contigs as well as full-length

cDNA sequences of C. elegans effectors were analysed using the NCBI Conserved Domain

Search Service (Marchler-Bauer et al. 2009, Marchler-Bauer et al. 2011) using default settings

and Pfam 27.0 programme (http://www.pfam.sanger.ac.uk) using protein sequence queries with

an e-value cut-off of 1.0 (default). In all cases, and to confirm that genomic contigs significantly

matched effectors, and in specific cases where e-values were higher than the threshold, the

tblastx search of the genomic contigs was conducted using sequences of specific functional

domains of C. elegans effectors. Multiple sequence alignments to compare matching contigs or

orthologues of effectors were done using MultAlin (Corpet 1988). To obtain open reading

frames from genomic contigs for any protein analysis, the Open Reading Frame Finder

(http://www.ncbi.nlm.nih.gov/projects/gorf) and the Translate programme at Expasy

(http://web.expasy.org/translate) were used. The exon prediction for parasitic nematode Dicer

genes was done using FGENESH (Solovyev et al. 2006) and tblastx against corresponding C.

40

elegans proteins, whilst graphics were created on the Exon-Intron Graphic Maker version 4 at

wormweb.org. Protein graphics were generated using protein sequences in Pfam 27.0.

2.3.4 Phylogenetic analyses

Multiple alignments for protein sequences using clustal W were created in MEGA version 6

(Tamura et al. 2013). Neighbour-joining trees with bootstrapping 1000 replicates were created

based on those alignments. The RdRp domain was used for analysis of RNA-dependent RNA

polymerase genes while PIWI domain was used for argonautes in all nematodes.

2.4 Results

2.4.1 Genomic contigs and ESTs of M. incognita, M. hapla, A. suum and B.

malayi with homologies to effectors of C. elegans

From the literature and database (wormbase.org version WS241) 87 genes were identified as

involved in the RNAi pathway of C. elegans, out of which two argonautes SAGO-2 (Wormbase

ID: WBGene00018921) and PPW-1 (Wormbase ID: WBGene00018921), were found to have

the same nucleotide sequence submitted in the database under different gene names.

‘Discontinued genes’ (rde-3, M03D4.6 and C06A1.4) were also omitted from the list and the

remaining 83 genes were divided into different groups based on their functions in the RNAi

pathway of C. elegans.

For the 83 selected effectors of C. elegans, the percentage of full length genes mapped to

genome contigs/scaffolds and ESTs of A. suum was 1.2% and 0.23% respectively. This

percentage was 0.66% and 0.053% for B. malayi, 3% and 0.31% for M. hapla and 1.69% and

0.34% for M incognita. Genes having homologues or coding for similar proteins within the

genome of C. elegans mapped significantly (10E-5) to a large number of contigs for each of the

four parasitic nematodes. These were the argonautes, spreading proteins haf-6 and sid-3, and the

genes encoding zinc finger domains i.e. zfp-1 and zfp-2. For M. incognita, two genes belonging

to different functional groups were present on the same contig (CABB01000055) i.e. tsn-1 and

mut-7. Some of the genes of PPNs e.g. pash-1, vig-1 and smg-6 were only identified after

alignment with animal parasitic nematode (APN) orthologues indicating that these genes were

more closely related than to those of C. elegans. The number of significantly mapped sequences

for each gene and the contig ID/Accession No. of the sequence coding protein domains for

specific effectors in parasitic nematodes are listed in supplementary Table 2.1 (Appendix).

41

2.4.2 Small RNA transport proteins

Four groups of these have been characterised in C. elegans, including those with systemic RNAi

defective phenotypes (sid-1, sid-2, sid-3, sid-5), those with RNAi spreading defective

phenotypes (rsd-2, rsd-3, rsd-6), exportins (xpo-1, xpo-2, xpo-3) and the haf-6 gene. A blastn,

blastx and tblastx analysis of the whole genome scaffold and contig sequences revealed that

three sequences of A. suum (AMPH01007595; ANBK01003199; AEUI01011716) and one of B.

malayi (AAQA01000131) had significant similarity to sid-1 of C. elegans. A further assessment

of these sequences indicated the presence of a type of sid-1 RNA channel similar to that present

in sid-1 of C. elegans. The amino acid sequence similarity is presented in Figure 2.1A. No ESTs

or genomic contigs of either M. incognita or M. hapla matched significantly to the sid-1 full

length gene sequence or the signature sequence of the SID-1 RNA protein. Also, no sequence

(ESTs or genomic contigs) for any other plant parasitic nematode at NCBI was significantly

similar to sid-1 sequence. Similarly, no contig of any of the four parasitic nematodes matched

significantly to the sid-2 gene of C. elegans. The C. elegans gene sid-3 encodes a protein

tyrosine kinase (PTK) domain together with an Src homology 3 (SH3) and a GTPase binding

domain responsible for efficiently importing dsRNA into cells (Jose et al. 2012). Protein

tyrosine kinases are the second largest family of proteins in C. elegans, with 411 identified

homologues (Plowman et al. 1999). It was therefore not surprising that a large number of

genomic contigs of all the four parasitic nematodes mapped significantly to this gene i.e. 100 for

A. suum, 59 for B. malayi, 26 for M. hapla and 30 for M. incognita. Protein domain analysis on

these contigs confirmed the absence of SH3 domain from all of those contigs. However, the

GTPase binding domain, together with the PTK domain, was found in three whole genome

sequences for A. suum (ANBK01007000; AMPH01017683; AEUI02000893), 2 for B. malayi

(AAQA01000282; CAPY01003672) and one each for each of the Meloidogyne spp.

(ABLG01000030; CABB01000892). As was the case for sid-2, no orthologues were found in

the genomic contigs of the four parasitic nematodes for sid-5.

Of the three RNAi spreading defective genes in C. elegans, sequences matching only rsd-3 were

identified for all four parasitic nematodes. Identified contigs contained the epsin amino-terminal

homology (ENTH) motif typical of the rsd-3 gene of C. elegans and cytosolic proteins of plants

which are involved in vesicle trafficking (Holstein and Oliviusson 2005). No orthologues of

rsd-2 and rsd-6 were identified in the EST and genomic contig databases for the four parasitic

nematodes considered in this study and also all other PPNs.

Contigs of A. suum, B. malayi, M. incognita and M. hapla with significant matches to the three

exportin genes of C. elegans were analysed for the presence of functional protein domains. They

were all identified in APNs but for the two RKN species only two contigs were found.

42

Functional domain analysis for M. hapla contigs ABLG01001363 and ABLG01000755

confirmed the presence of xpo-1 and xpo-2 as the two contigs had all the protein domains for

these two genes, while M. incognita contig CABB01000462 coded for xpo-2. Interestingly,

detailed analysis revealed that when two M. incognita contigs, CABB01002745 and

CABB01004119 were re-assembled, the super contig matched completely to the full length C.

elegans xpo-1 mRNA (Figure 2.1B). There was no contig found coding for the xpo-3 gene in

the two PPNs.

(A)

(B)

The haf-6 gene is a member of the ATP binding cassette (ABC) transporter gene family, and the

C. elegans genome encodes 60 proteins with ABC transporter domains (Sheps et al. 2004).

Unsurprisingly, a large number of contigs of A. suum (95), B. malayi (32), M. hapla (14) and M.

incognita (17) mapped to this gene. Parasitic nematodes such as A. suum, B. malayi and

C. elegans Xpo1

C. elegans Xpo2

C. elegans Xpo3

Mi (CABB01004119 + CABB01002745)

Mi (CABB01000462)

Mh (ABLG01001363)

Mh (ABLG01000755)

Figure 2.1: (A) Conserved signature amino acids of sid-1 RNA channel in C. elegans with

mapped contigs of A. suum (As) and B. malayi (Bm). (B) Protein domain architecture of the

three exportins of C. elegans (Ce) compared to two contigs each of the M. incognita (Mi) and

M. hapla (Mh) revealing that they are xpo-1 and xpo-2.

43

Meloidogyne spp. are known to be susceptible to environmental RNAi, and although there are

degrees of susceptibility, they seem to have a smaller but similar repertoire of genes involved in

dsRNA uptake and spread.

2.4.3 The Dicer and associated genes

The Dicer gene plays a central role in RNAi pathways of eukaryotes. For organisms which

encode only one dicer, it is responsible for processing all forms of dsRNAs into small RNAs,

including siRNAs and miRNAs. The Dicer-1 of C. elegans was used to identify, characterise

and compare the structures of dicers in the parasitic nematodes. The lengths of the predicted

pre-mRNA for both M. incognita and M. hapla dcr-1 are similar to that of C. elegans although

they have slightly more exons (Figure 2.2A).

Genomic contigs with the most sequence identity to C. elegans dcr-1 were CABB01000157

(42843 bp) for M. incognita, ABLG01001138 (50884 bp) for M. hapla, CAPY01005536

(830985 bp) for B. malayi and ANBK01006853 (13709 bp), AMPH01008524 (10740 bp) and

AEUI02001038 (51650 bp) for A. suum. The drh-1 and drh-3 genes of C. elegans have three

similar functional domains each i.e. DEXDc (DEAD-like helicase), HELICc (Helicase C-

terminal domain) and RIG-1_C-RD (C-terminal domain of RIG-1). Although the mRNA

sequences of drh-1 and drh-3 were significantly identical to contigs of the four parasitic

nematodes, functional analysis revealed striking differences in the protein domains of these

genes. Whereas these contigs had the DEXDc and HELICc domains, the RIG-1 domain was

conspicuously missing in contigs matching to drh-1 of C. elegans. In drh-1 and drh-3 of C.

elegans, there is only 24% identity between the RIG domains. A detailed analysis of whole

genome contigs of the parasitic nematodes as well as ESTs and with those of H. glycines, B.

xylophilis and H. contortus indicated that this domain is absent in available sequences of

parasitic nematodes.

Unlike rde-4 which encodes two dsRNA binding motifs and did not match significantly to

whole genome contigs of any of the four parasitic nematodes, contigs mapping to C. elegans

argonaute rde-1, were 14 for A. suum, 12 for B. malayi and four each for M. hapla and M.

incognita: nematodes have a suite of argonaute proteins as important players of gene silencing.

The only functional domain of pir-1 that is ‘Dual specificity phosphatase catalytic domain’, was

identified in six contigs of A. suum, six for B. malayi, two for M. hapla and three for M.

incognita.

44

D. melanogaster DCR1

D. melanogaster DCR2

A. thaliana DCL1

A. thaliana DCL2

A. thaliana DCL3

A. thaliana DCL4

(A)

(B)

(C)

Nematode Exons Unspliced

Gene (bp)

C. elegans dcr-1

27

8,420

M. hapla dcr-1

34

8,465

M. incognita dcr-1

35

8,393

B. malayi dcr-1

40

19,640

A. suum dcr-1

40

32,216

Figure 2.2: (A) Graphical representation of dcr-1 gene of C. elegans compared to the four

parasitic nematodes with gaps indicating intron regions (Scale=1000 bp). The seven protein

domains are DEXDc. Helicase C-Terminal (Helicase CT), Dicer Dimer, PAZ, two Ribonuclease

III C-terminal domains (Ribo III-CT) and double stranded RNA binding motif (DSRM).Whole

genomic contigs mapped to C. elegans dcr-1 were used to predict protein domain coding regions

(coloured) in the contigs of M. hapla (ABLG01001138), M. incognita (CABB01000157), B.

malayi (CAPY01005536) and A. suum (AEUI02001038; AMPH01008524; ANBK01006853).

Number of exons and the unspliced sequence length (bp) are also indicated. (B) Drosophila

melanogaster protein domain architecture for the two Dicers. (C) Differences in the protein

domain architecture between the four Arabidopsis thaliana Dicer-like (DCL) gene products.

45

The best matching contigs of A. suum (AEUI02000028), B. malayi (AAQA01000005), M. hapla

(ABLG01000521) and M. incognita (CABB01000477) to the mRNA sequence of C. elegans

Drosha (drsh-1) encode the drsh-1 specific protein domains; two Ribonuclease III C terminal

domains and a double stranded RNA binding motif (DSRM). The closest matches for the drsh-1

cofactor Pasha (pash-1) in the four parasitic nematodes had low total bit scores and coverages:

154 (15%) for A. suum, 155 (12%) for B. malayi, 161 (15%) for M. hapla and 161 (15%) for M.

incognita contigs. However, when the characterized pash-1 gene of A. suum (HQ611976) was

used as query to identify this gene in B. malayi, M. hapla and M. incognita, it mapped to the

same contigs but with better identity scores indicating they are more closely related to each

other than to C. elegans.

2.4.4 RNA-induced silencing complex (RISC)

The four known proteins that interact to form the RISC complex in C. elegans are ain-1, ain-2,

tsn-1 and vig-1. Both of the alg-1 interacting proteins ain-1 and ain-2 encode an M domain (M

domain of the protein GW182). Apart from B. malayi contig (AAQA01000363) coding for ain-

1, there were no significant matches for these two genes in any contigs or ESTs of the four

parasitic nematodes when the ain-1 coding sequence of B. malayi was used as query to search

for matches in A. suum, M. hapla and M. incognita. Significant matches were identified only for

contigs of A. suum but protein domain analysis revealed that those sequences did not code for

the M domain.

The C. elegans tsn-1 gene encodes six functional protein domains (Figure 2.3); four SNase

(Staphylococcal nuclease) domains arranged in tandem followed by a Tudor domain fused with

a fifth SNase domain at the C-terminus. These domains are not significantly identical to each

other: the second and fourth SNase domains have the highest total score of only 27%. Contigs

with high identities to the mRNA sequence of tsn-1 of C. elegans are present amongst contigs of

all the four parasitic nematodes. Protein domain analysis revealed the presence of both types of

functional domains in all the contigs but with a lower identity score for the Tudor domain of M.

hapla and M. incognita compared to that of C. elegans. There seems to be structural differences

in proteins of the different nematodes where C. elegans and M. incognita have five SNase

domains all seemingly arranged similarly. But all contigs analysed for M. hapla, B. malayi and

A. suum indicate the presence of only the four SNase domains, each missing particularly the

fifth domain fused to the Tudor domain which is present in C. elegans and M. incognita (Figure

2.3A).

The functional significance of these differences, if indeed it is the case, is not known and like

many other differences, need to be investigated further. Interestingly, the M. incognita whole

46

genome contig CABB01000055 is 63,544 bp long and encodes the functional domains of the

tsn-1 gene as well as mut-7, nuclear RNAi effector, which in C. elegans is located on a different

chromosome.

The vig-1 gene was identified in whole genome sequences of A. suum (AMPH01002339) and B.

malayi (AAQA01001685), but not for PPNs. However, when the identified vig-1 domain of this

B. malayi contig was used as a query, contig ABLG01000254 of M. hapla and contig

CABB01000081 of M. incognita were identified, but with very low identity scores. Detailed

analysis of these contigs confirmed the presence of the mRNA binding protein domain

HABP4_PAI-RBP1 similar to that of C. elegans and the animal parasitic nematodes

Schistosoma japonicum (Q5DA16) and Schistosoma mansoni (Q9N2M6).

2.4.5 RNAi amplification

Seven genes encoding proteins involved in the siRNA amplification process have been

described in C. elegans. These are ego-1, rrf-1, smg-2, smg-5, smg-6, rde-10 and rde-11. The

genes ego-1 and rrf-1 of C. elegans have only one RdRp domain (77% nucleotide similarity and

59% amino acid identity between them) are both present on chromosome I, only 0.9 kb apart

(Sijen et al. 2001). Consequently the same genomic contigs of the parasitic nematodes matched

significantly to both C. elegans rrf-1 and ego-1 (total bit scores between 497-1760); three each

for A. suum and B. malayi, two for M. hapla and four for M. incognita. All of these sequences

had the RdRp domain with the signature motif ‘DbDGD’. The full length mRNA sequences of

both genes were covered in most of contigs and scaffolds of the parasitic nematodes.

Contigs encoding genes similar to the smg-2 were found in sequences of all the four parasitic

nematodes and contained all three functional domains (AAA_30, AAA_12 and UPF1_Zn_bind)

typical of the C. elegans smg-2 gene. The C. elegans smg-6 matched with significant identities

to A. suum and B. malayi sequences. The best matching contigs for M. hapla (ABLG01000285)

C. elegans TSN1

M. incognita TSN1

M. hapla TSN1

A. suum TSN1

B. malayi TSN1

Figure 2.3: Protein domain analysis of nematode TSN1 indicating the missing

Staphylococcal nuclease (SNase) domain after the Tudor domain in M. hapla, A. suum and B.

malayi.

47

and M. incognita (CABB01000011) had insignificant e-values of 0.029 and 0.012 respectively.

However, when smg-6 coding sequence of B. malayi was used as a query, significant matches

were identified in contigs of M. incognita and M. hapla. These had the signature functional

domains present in C. elegans smg-6 namely EST1, EST1 DNA binding domain and PINsmg6

domain. No genomic contig, scaffold or EST of any of the parasitic nematodes was similar or

identical to the full length mRNA sequences of C. elegans smg-5 (or its endcoded domain PIN

smg5) or two other genes involved in RNAi amplification, rde-10 and rde-11.

2.4.6 RNAi inhibitors

Although their function is not exclusive to inhibition of RNAi, 13 such genes for which loss of

function increases the sensitivity of RNAi, have been investigated in C. elegans. These are

enhanced RNAi phenotype genes (eri-1, eri-3, eri-5, eri-6/7, eri-9), adenosine deaminase acting

on RNA genes (adr-1, adr-2), XRN (mouse/S. cerevisiae) ribonuclease related genes (xrn-1,

xrn-2), the RdRp containing genes rrf-3, lin15-b, gfl-1 and zfp-2.

The eri-1 gene in C. elegans which removes the 2 nt overhangs at 3′ ends of siRNAs thereby

restricting uptake by the RISC, has two functional domains i.e. SAP which is a putative

DNA/RNA binding domain and ERI-1, a DEDDh 3′-5′ exonuclease domain. Spliced mRNA

sequence of eri-1 mapped significantly to three contigs each for A. suum and B. malayi, one of

M. hapla and two of M. incognita. Also it appeared that contigs of B. malayi, M. hapla and M.

incognita do not encode the SAP domains (Figure 2.4A). Whether this represents a functional

difference or divergence in the gene is not known.

Only one and two contigs respectively for A. suum and B. malayi were significantly identical to

eri-5 of C. elegans: all encoded the Tudor domain typical of this gene. The tudor domain of eri-

5 is completely different in terms of amino acid sequence from the Tudor domain of ekl-1 which

is a nuclear RNAi effector and tsn-1 gene of the RISC complex. However, no contigs or ESTs

of M. hapla or M. incognita were found to encode a Tudor protein domain. In C. elegans, the

eri-6/7 mRNA is formed by trans-splicing of two pre-mRNAs eri-6 and eri-7 (Fischer et al.

2008) and encodes two P loop motifs designated as AAA_11 and AAA_12. These P loop motifs

are also present in the smg-2 gene designated as AAA_30 and AAA_12 but have different

amino acid sequences than the eri-6/7 domains. The eri-6/7 mRNA mapped significantly to A.

suum and B. malayi contigs. Out of the 83 effectors of RNAi analysed in this study, the eri-6/7

gene was the only gene not present in both Meloidogyne spp., although it was in M. hapla

(ABLG01001054). The genes eri-3 and eri-9 of C. elegans had no significant matches to any of

the four parasitic nematodes.

48

The rrf-3 gene has been reported as an inhibitor of RNAi in C. elegans and rrf-3 mutants with

enhanced sensitivity to RNAi have been used in several experiments to assess effects of

knockdown of several genes. One hypothesis is that it competes with ego-1 and rrf-1 for

components of the RNAi machinery (Simmer et al. 2002). Although C. elegans rrf-3 is quite

different from ego-1 and rrf-1 (58% and 57% identity respectively), the same contigs of the

parasitic nematodes (four for M. incognita, two for M. hapla, ten for A. suum and seven for B.

malayi) matched to all three genes. This must be because both the genes and the contigs code

for RdRp domains. To explore the identity of the RdRp domains of these genes and the contigs,

a phylogenetic tree was constructed using translated amino acid sequences of the domains.

Contigs coding for 100% identical RdRp domains were excluded, and RdRp domains of the

same genes in C. brigssae and C. ramenei were included in the analysis (Figure 2.4B). The tree

indicates close relationships of ego-1 and rrf-1 both of which are distantly related to rrf-3. One

RdRp each for A. suum (ANBK01005062) and B. malayi (CAPY01003132) clustered with rrf-3

of C. elegans whereas two each for these species clustered with the rrf-1 and ego-1 RdRps of

Caenorhabditis spp. It appears the RdRps of the PPNs are distantly related to those of the free

living nematodes. While it may be safe to suggest the presence of ego-1 and rrf-1 in the genome

of the APNs, it was not possible to distinguish the RdRps of the PPNs.

The effector gfl-1 was conserved in all four parasitic nematodes. The genes xrn-1 and xrn-2 are

5′-3′ exoribonucleases similar in their protein functional domain architecture. They code for the

same Xrn_N domains which in C. elegans has 52% similar amino acids. Both of these inhibitor

sequences mapped to significantly matching contigs of four parasitic nematodes. Two contigs

for M. incognita (CABB01001503; CABB01003205) mapped to different parts of the C.

elegans xrn-2 completing the full length gene. Further analysis suggested that APNs have both

of these RNAi inhibitors but PPNs possess one of them, which based on the e-value and total

scores was xrn-2.

The adr-1 gene of C. elegans has three functional domains i.e. two DSRM and one Adenosine-

deaminase (A_deamin) domain while adr-2 has one DSRM and one A_deamin domain which

suppresses RNAi by deaminating transgenic dsRNA. A. suum contig (AMPH01003945) that

mapped to adr-1 codes for one DSRM domain with low identity to C. elegans DSRM while the

whole genome scaffold AMPH01015223 mapped for adr-2 codes for the A_deamin domain

only. These results were similar for B. malayi contigs mapping to adr-1 and adr-2. There was

no significant match of any contig of Meloidogyne spp. to the C. elegans adr-1 or adr-2 (Figure

2.4C).

49

(A)

(B)

(C)

Figure 2.4: (A) Protein domain composition of the RNAi inhibitor ERI1 of C. elegans

compared to that of parasitic nematodes. (B) Phylogenetic analysis of the RNA-dependent

RNA-polymerase genes of C. elegans, A. suum (As), B. malayi (Bm), M. hapla (Mh) and M.

incognita (Mi). The Neighbour-joining tree was constructed based on ClustalW alignment of

RdRp domain sequences. (C) Protein domain architecture of the inhibitor gene adr-1 and adr-2

of C. elegans compared to that of the two animal parasitic nematodes.

50

C. elegans mutants of lin-15b are more sensitive to RNAi. This gene encodes a protein that

contains the THAP functional domain, which is a putative DNA-binding domain. Neither the

gene sequence nor the THAP domain matched significantly to any contig or EST of the four

parasitic nematodes. The full length mRNA of C. elegans zfp-2 mapped significantly to 40

genomic contigs of A. suum, 100 contigs of B. malayi, 17 of M. hapla and 20 of M. incognita.

The reason for this large number of matching contigs is that the zinc finger proteins are amongst

the most abundant proteins in eukaryotic genomes and are needed for stabilising other protein

structures and as part of several transcriptional factors (Haerty et al. 2008).

2.4.7 Nuclear RNAi effectors

Expressed in both the cytosol and the nucleus, 17 C. elegans genes are known to play vital roles

in the nuclear RNAi process. Eight of these were not identified from genomic contigs or ESTs

of any of the parasitic nematodes. These were the nuclear RNAi defective genes (nrde-1, nrde-2

and nrde-4), rde-2, the mutator phenotypic genes (mut-2, mut-7), mes-3 and two enhancers of

ksr-1 lethality (ekl-1 and ekl-5). Three contigs for A. suum and two for B. malayi and one each

for M. hapla and M. incognita mapped to C. elegans ekl-4 and they all encode for the gene

specific DMAP1 functional domain. For ekl-6 gene, two contigs each for the animal parasitic

nematodes mapped to C. elegans ekl-6 and encode for the DUF2435 protein domain. This gene

was not found in the contigs of PPNs.

The gene mut-7 was identified in contigs of all the four parasitic nematodes. The domain mut-7

characteristic of the gene was less identical to the one identified in contig CABB01000055 for

M. incognita. For the six effectors mut-2, mes-2, mes-6, cid-1, rha-1 and zfp-1, significantly

matching contigs of the parasitic nematodes contained all the functional domains typical of

these genes. Nuclear RNAi defective (nrde) genes in C. elegans have been associated with

heritable RNAi. Interestingly, there is no detailed study on heritable RNAi in parasitic

nematodes although there are suggestions of persistence of gene knockdown in some percentage

of root-knot nematodes in some studies.

2.4.8 Argonautes

Twenty-eight argonautes which function at different stages of the RNAi and miRNA pathways

in C. elegans have been reported (Yigit et al. 2006). Since then rde-3 has been renamed mut-2, a

nuclear RNAi effector and M03D4.6 and C06A1.4 have been designated as pseudogenes: these

were excluded from the analysis. Also during the analysis, it was found that ppw-1 and sago-2

sequences were identical, and so one was excluded leaving 24 argonaute sequences for study.

These were analysed for the presence of functional protein domains: five of them (alg-1, alg-2,

51

alg-4/tag-76, T22B3.2, T23D8.7) encode PAZ, PIWI and another domain known as DUF1785.

The C04F12.1, ZK218.8 and ZK1248.7 genes encode only the PIWI domain while the rest

(R06C7.1, F58G1.1, rde-1, C16C10.3, ppw-1/sago-2, ppw-2, sago-1, csr-1, T22H9.3, ergo-1,

prg-1, prg-2, F55A12.1, nrde-3, Y49F6A.1, C14B1.7) code for both PAZ and PIWI domains.

Using full-length sequences of the C. elegans argonautes, a total of 13, 10, 13 and 16 contigs

respectively for A. suum, B. malayi, M. hapla and M. incognita were identified with significant

alignment scores. This means 11 unique sequences for A. suum, six for B. malayi, ten for M.

hapla and 11 for M. incognita were used to analyse the presence of PAZ/PIWI domains and for

constructing phylogenetic tree with argonautes of C. elegans. As with the C. elegans genes, all

the contigs of parasitic nematodes have the conserved PIWI domain. The cladogram shows the

phylogenetic relationship between C. elegans and both parasitic groups of nematodes based on

the PIWI domain (Figure 2.5).

Based on sequence similarity, there are distinctly different sub clades for example, in clade 1,

where alg-1, alg-2 and T 23D8.7 are relatively similar to sequences of each of the parasitic

nematodes. The PIWI domains of C. elegans argonautes appear to be distantly related to those

of the four parasitic nematodes. In both clades 2 and 6, generally, sequences of the parasitic

nematodes appear more closely related to each other than those of C. elegans. There appears to

be a clear distinction in clade 3 where no argonaute of parasitic nematodes clustered with the

five C. elegans argonautes (F58G1.1, PPW2, ZK1248.7, R06C7.1, F55A12.1) that are involved

in transcriptional silencing and germline RNAi while only A. suum PIWI coded by the contig

AMPH01002307 displayed homology to the argonaute ERGO1 in the sub clade 7. Argonautes

involved in nuclear RNAi (nrde-3, C16C10.3, T22H9.3, Y49F6A.1) define a clade on their own

but do appear to be distantly homologous to one argonaute each for the four parasitic

nematodes.

52

Figure 2.5: Neighbour-Joining cladogram based on the conserved PIWI domain of all

argonaute sequences in C. elegans and four parasitic nematodes A. suum (As). B. malayi (Bm),

M. hapla (Mh) and M. incognita (Mi). Contig IDs of the sequences coding for the argonaute

proteins are indicated in parenthesis.

2.5 Discussion

The aim of the work presented in this chapter was to study the various components of the sRNA

pathways of parasitic nematodes, to explore the similarities and differences that exist when

compared to the model nematode C. elegans. The analysis shows that out of the 83 genes

described in the RNAi and miRNA pathway of C. elegans, only 49 homologous sequences were

found in the available genomic contigs of A. suum, 45 in B. malayi contigs, and 40 each in M.

hapla and M. incognita contigs, with the majority displaying conserved protein domain

architecture. Protein domain similarity and identity scores for these genes indicate that PPNs are

more similar to APNs than to C. elegans, only in relation to the whole genome sequences

analysed for RNAi pathway genes (Appendix-Supplementary Table 2.2).

1 6

2

4

3

7

5

53

Among the sequences analysed, the Dicer-1 was the most conserved protein, with its seven

domains present in all the nematodes studied. All eukaryotes displaying RNAi ability have

Dicer-like genes involved, with four classes in Arabidopsis, two in insects and fungi and one

characterised for mouse and humans. Homology studies identified differences in the functional

domains of Dicer-like genes of the five nematodes compared to those of other organisms. There

were differences in the length of the genomic sequences and the number of exons coding for the

protein. These suggest structural differences in the dicers even though they appear to encode

similar protein domains. It is possible that the distance between the PAZ and ribonuclease III

domains of the dicer protein, which is a measure of the size of siRNAs produced (MacRae et al.

2006), may be responsible for the different sizes of siRNAs generated by C. elegans (~22-23 bp,

Ketting et al. 2001) and those produced by plants (21, 22 and 24 bp for Arabidopsis). The

length of siRNAs produced by parasitic nematodes has not been investigated yet and the

identification and characterisation of sequences potentially encoding the dicer of PPNs such as

undertaken in this study, is the first step for characterising in detail the functions of dicer in

development in PPNs.

The C. elegans dicer has been associated with rde-1 and rde-4, the mutants of which

demonstrate a complete absence of RNAi response to foreign dsRNA (Tabara et al. 1999;

Tabara et al. 2002). However, these two essential genes were absent from all of the parasitic

nematodes. This indicates a possible alternative mechanism for dsRNA retention in these

nematodes for example, in Drosophila, RISC contains DCR-2 protein as part of the complex

and appears to play a role in retaining the dsRNA that enters the cell thereby maintaining an

unprocessed pool of dsRNA (Kim et al. 2007; Shih and Hunter 2011).

In plants, insects and C. elegans, the RNAi pathway delivers robust antiviral response. In C.

elegans the DRH-1 protein has a RIG-I domain which senses and binds viral single-stranded

RNA (Lu et al. 2009). This domain in vertebrates also binds to viral RNA bearing 5′-phosphates

triggering antiviral responses (Rehwinkel et al. 2010). The parasitic nematodes studied here all

lacked a detectable RIG-I domain in the drh-1 sequence. Mutation in the drh-1 in C. elegans

leads to the loss of antiviral response and it is suggested that this protein may act at the top of

the antiviral response cascade triggering an immune response (Ashe et al. 2013). This disparity

points towards an alternate mechanism for recognition of viral RNA and resulting RNAi-based

antiviral response mechanisms for this group of nematodes. Mutants of rde-1 and rde-4 in C.

elegans are more susceptible to virus replication while rrf-3 and eri-1 mutants have increased

antiviral response (Lu et al. 2005; Wilkins et al. 2005).

The orthologues of rde-1, rde-4 and rrf-3 which are involved in the antiviral defence

mechanism appear to be missing in the genomic sequences of all four parasitic nematodes.

54

These nematodes also lacked the SAP domain in the sequence coding for the gene eri-1. Yeast

assays show that the SAP domain is responsible for binding dsRNA and that ERI-1 or the SAP

domain when expressed separately do not result in dsRNA degradation (Iida et al. 2006). This

perplexing difference in the proteins affecting antiviral response in these parasitic nematodes

requires further investigation to explore their role in this pathway if any. Another possible

explanation could be that the parasitic nematodes, specifically those included in this study, live

in a confined environment i.e. inside a host and so are less exposed to hostile environments,

possibly resulting in a loss of robust antiviral defences.

Comparative analysis also showed differences for RNAi spreading genes between C. elegans

and parasitic nematodes. For PPNs, the absence of sid-1, sid-2 and other spreading proteins

seem to have no effect on dsRNA uptake by soaking which when absent in C. elegans, impairs

RNAi. This indicates that other mechanisms or unidentified genes may also be involved in this

process. It is also possible that sid-3, rsd-3 and haf-6 proteins, which are present in both of these

plant parasitic nematodes, have evolved to play the role of spreading dsRNA through tissues,

although SID-3 lacks two of the protein domains SH3 and GTPase which are involved in signal

transduction and protein binding. Effectors of the miRNA pathway seem to be more conserved

even though ain-1 and ain-2 were not found in any of the parasitic nematodes except B. malayi

in which ain-1 was present. These two proteins of the RISC machinery seem to be specific to

C. elegans only. Immunoprecipitation and microarray analysis indicates the association of these

two RISC proteins exclusively with miRNAs (Zhang et al. 2007).

In C. elegans, RNAi has been shown to persist in the next progeny, and this effect is proposed

to be dependent on amplification of a trigger (Alcazar et al. 2008). This germline amplification

effector (ego-1) is present in all the four parasitic nematodes, but the nuclear RNAi effectors

nrde-1, nrde-2 and nrde-4, which are the core components carrying out heritable RNAi in C.

elegans, were not found. The nrde-3 gene, which encodes the argonaute responsible for

localising secondary siRNAs to the nucleus, was also one of the argonautes that had low

significant matches to the argonautes of parasitic nematodes. This suggests a large divergence or

the complete absence of this effector and the possibility of absence of heritable RNAi in these

species, although this aspect has not been investigated experimentally.

Argonaute proteins show a striking contrast in numbers, with 24 in C. elegans and fewer than

half of these proteins in the parasitic nematodes. Overall there were fewer argonaute proteins

and less conservation in nuclear, germline and transcription related argonautes, which suggests

differences in these mechanisms/biological processes. It is possible that argonautes of parasitic

nematodes act as convergent points for the various processes involved, or carry out more than

one function in the sRNA pathways with the aid of other associated proteins.

55

Table 2.1 summarises the small RNA effector repertoire in C. elegans compared to the four

parasitic nematodes studied here with already published data of Pratylenchus coffeae,

Globodera pallida and Bursaphelenchus xylophilus. Despite their morphological similarities,

and more than 40% highly homologous genes to C. elegans, parasitic nematodes have a

completely different life style (Hashmi et al. 2001). The remaining 60% of the predicted genes

have probably diverged more when compared to C. elegans, or belong to a completely different

set reflecting their parasitic nature, and the requirement to counteract host defence mechanisms

for successful parasitism. The presence of eri-6/7 in M. hapla in the genome sequences and

absence of the same in sequences of the closely related M. incognita is striking if infact this is

the case, since RNAi is a conserved mechanism. Phylogenetically, M. incognita and M. hapla

belong to different clades and have different modes of reproduction (De Ley et al. 2002). M.

incognita reproduces by mitotic parthenogenesis whereas M. hapla can exhibit both mitotic and

meiotic parthenogenetic modes of reproduction (Bert et al. 2011). Since they have a more

complicated lifestyle with different moults and developmental phases, the endo-RNAi and

miRNA pathways of parasitic nematodes may be more sophisticated as was initially proposed,

when compared to that of C. elegans.

The disparities between the RNAi effectors of C. elegans and the four parasitic nematodes

studied here indicate wide differences between these nematodes at the sRNA level. Parasitic

nematodes such as PPNs can affect host gene expression, but the precise mechanisms involved

are still under intense research. The absence of 34 to 43 RNAi effectors in the genomic

sequences of parasitic nematodes compared to those present in C. elegans requires explanation

through further functional studies. A host of argonautes performing different functions in the

small RNA pathways of C. elegans, but missing in parasitic nematodes, may indicate a more

complicated mechanism and diversity of effector functions for these nematodes, and also the

possibility of involvement of as yet unidentified genes. On an evolutionary basis, C. elegans

may show high conservation of all members of a protein family with the parasitic nematodes,

but it is possible that the parasitic nematodes have more conserved protein families among

them. Another possibility for absence of sRNA pathway genes in parasitic nematodes may be

that the sequences have diverged so much that it is not possible to identify them using only

sequence-homology-based programmes.

56

Table 2.1: Comparison of the RNAi pathway genes of four parasitic nematodes with the published data

of G. pallida, B. xylophilus and P. coffeae (Cotton et al. 2014; Kikuchi et al. 2011; Burke et al. 2015).

C. elegans A. suum B. malayi M. hapla M. incognita P. coffeae G. pallida B. xylophilus

Transport proteins

xpo-1, xpo-2,

xpo-3, rsd-2,

rsd-3, rsd-6,

sid-1, sid-2,

sid-3gpb, sid-

5gpb, haf-6gpb

xpo-1, xpo-

2, xpo-3,

rsd-3, sid-1,

sid-3, haf-6

xpo-1, xpo-

2, xpo-3,

rsd-3, sid-1,

sid-3, haf-6

xpo-1, xpo-2,

rsd-2, rsd-3,

sid-3, haf-6

xpo-1, xpo-2,

rsd-2, rsd-3,

sid-3, haf-6

xpo-1, xpo-

2, rsd-3

xpo-1, xpo-2,

rsd-3,

xpo-1, xpo-2,

rsd-6

Dicer and associated proteins

dcr-1, drh-1,

drh-3, pir-1gpb,

drsh-1, pash-

1, rde-4

dcr-1, drh-1,

drh-3, pir-1,

drsh-1,

pash-1

dcr-1, drh-1,

drh-3, pir-1,

drsh-1,

pash-1

dcr-1, drh-1,

drh-3, pir-1,

drsh-1, pash-1

dcr-1, drh-1,

drh-3, pir-1,

drsh-1, pash-1

dcr-1, drh-3,

drsh-1

dcr-1, drh-3,

drsh-1, pash-

1

dcr-1, drh-1,

drh-3, drsh-1,

pash-1, rde-4

RNA Induced Silencing Complex (RISC)

ain-1, ain-2,

tsn-1, vig-1

tsn-1, vig-1 tsn-1, vig-1,

ain-1,

tsn-1, vig-1 tsn-1, vig-1 tsn-1 tsn-1, vig-1,

ain-1

RNAi Amplification

ego-1, rrf-1,

smg-2, smg-5,

smg-6, rde-

10gpb, rde-

11gpb

ego-1, rrf-1,

smg-2, smg-

6

ego-1, rrf-1,

smg-2, smg-

6

ego-1, rrf-1,

smg-2, smg-6

ego-1, rrf-1,

smg-2, smg-6

ego-1, rrf-1,

smg-2, smg-

6

ego-1, smg-

2, smg-6,

ego-1, rrf-1,

smg-2, smg-6

RNAi Inhibitors

eri-1, eri-3,

eri-5, eri-6/7,

eri-9gpb, adr-1,

adr-2, xrn-1,

xrn-2, rrf-3,

lin15-b, gfl-1,

zfp-2gpb, somi-

1gp

eri-1, eri-5,

eri-6/7, adr-

1, adr-2,

xrn-1, xrn-2,

xrn-3, gfl-1,

zfp-2

eri-1, eri-5,

eri-6/7, adr-

1, adr-2,

xrn-1, xrn-2,

xrn-3, gfl-1,

zfp-2

eri-1, eri-6/7,

xrn-2, gfl-1,

zfp-2

eri-1, xrn-2,

gfl-1, zfp-2

eri-6/7, rrf-

3, gfl-1

eri-1, xrn-2,

gfl-1

Adr-1, xrn-2,

rrf-3, gfl-1,

somi-1

Nuclear RNAi effectors

nrde-1gpb,

nrde-2 gpb,

nrde-4 gpb, rde-

2, mut-2p, mut-

7, mes-2, mes-

3, mes-6p, ekl-

1, ekl-4, ekl-5,

ekl-6, cid-1,

rha-1, zfp-1g

mut-2, mut-

7, mes-2,

mes-6, ekl-4,

ekl-6, cid-1,

rha-1, zfp-1

mut-2, mut-

7, mes-2,

mes-6, ekl-4,

ekl-6, cid-1,

rha-1, zfp-1

mut-2, mut-7,

mes-2, mes-6,

ekl-4, cid-1,

rha-1, zfp-1

mut-2, mut-7,

mes-2, mes-6,

ekl-4, cid-1,

rha-1, zfp-1

cid-1, ekl-1,

ekl-4, mes-2,

rha-1

cid-1, mes-2,

ekl-4, rha-1

Mut-7, mes-2,

mes-6, ekl-1,

ekl-4, ekl-6,

cid-1, rha-1

Argonautes

alg-1, alg-2,

alg-4/tag-76g,

T22B3.2b,

T23D8.7,

R06C7.1,

F58G1.1, rde-

1, C16C10.3,

ppw-1/sago-2,

ppw-2, sago-1,

csr-1,

T22H9.3,

ergo-1, prg-1,

prg-2,

F55A12.1,

nrde-3,

Y49F6A.1,

C14B1.7gb,

C04F12.1,

ZK218.8gp,

ZK1248.7

alg-1, alg-2,

T22B3.2,

R06C7.1,

rde-1, ergo-

1, F55A12.1,

Y49F6A.1,

C04F12.1,

ZK1248.7

alg-1, alg-

4/tag-76,

R06C7.1,

C16C10.3,

ergo-1,

C04F12.1,

alg-1, alg-

4/tag-76,

R06C7.1,

F58G1.1, csr-

1, T22H9.3,

F55A12.1,

Y49F6A.1,

C04F12.1

alg-1, alg-

4/tag-76,

R06C7.1,

F58G1.1,

C16C10.3,

F55A12.1,

C14B1.7,

C04F12.1,

ZK1248.7

alg-1, alg-2,

alg-4/tag-

76, T22B3.2,

T23D8.7,

R06C7.1,

F58G1.1,

rde-1,

C16C10.3,

ppw-1/sago-

2, ppw-2,

sago-1, csr-

1, ergo-1,

prg-1, prg-2,

F55A12.1,

nrde-3,

Y49F6A.1,

C04F12.1,

ZK1248.7

alg-1,

R06C7.1,

F58G1.1,

C16C10.3,

T22H9.3,

F55A12.1,

Y49F6A.1,

ZK1248.7

alg-1, alg-2,

T23D8.7,

R06C7.1, rde-

1, csr-1,

Y49F6A.1,

ZK1248.7

g = not determined in G. pallida, p = not determined in P. coffeae, b = not determined in B. xylophilus

57

Chapter 3

Identification of target genes from among sRNA

pathway effectors of M. incognita for nematode control

via in vitro RNAi

58

Identification of target genes from among sRNA pathway effectors of M.

incognita for nematode control via in vitro RNAi




3.1 Abstract

At present it is not possible to mutagenise or transform plant parasitic nematodes, therefore

functional analysis of their genes involves using RNAi, that is, delivering dsRNA/siRNA

complementary to the gene to be studied, either by in vitro soaking or via transgenic plants.

Previous studies using RNAi to probe gene function have mainly focussed on study of genes

involved in parasitism and development. In this study, 27 genes involved in the RNAi pathway

of the root-knot nematode M. incognita were subjected to in vitro RNAi assays, and their

subsequent infectivity and development was investigated after inoculation to tomato plants. In

response to soaking with dsRNA, J2s of M. incognita exhibited abnormal phenotypes after

RNAi of 25 of the genes tested: abnormal phenotypes included body curling, wavy body

movements and paralysis. On infection of tomato plants, RNAi of 12 of the genes reduced

nematode infectivity by ≥ 50%. The greatest reduction in infection (90%) was observed for

nematodes fed with dsRNA targeting dcr-1. Additionally, RNAi treatment of nematodes did

affect their development, manifested after moults by the development to adult females:

symptoms included elongated and transparent bodies compared to the normal saccate white

adult females. The results of screening target genes by soaking, and analysing the effects on

phenotype, and the longer term effects on nematode reproduction and development after

soaking and plant infection, confirm that in vitro screening is a valid approach to identify target

genes likely to be most effective in conferring nematode resistance by reducing infection and

affecting their normal development.

Keywords: In vitro RNAi, dsRNA soaking, M. incognita, RNAi pathway, nematode infection.

59

3.2 Introduction

Experiments in which gene knockdown in C. elegans was achieved via introduction of

complementary synthetic dsRNAs revolutionised nematode functional genomics (Fire et al.

1998). This strategy has since been used to study gene function in a wide range of organisms

including plants, fungi and insects (Chuang and Meyerowitz 2000; Khatri and Rajam 2007;

Tomoyasu et al. 2008). The recent availability of transcriptomic and genomic data for parasitic

nematodes has also opened doors for more extensive genomic studies of these organisms.

Unlike C. elegans, the difficulty in generating mutants, and lack of a routine method to generate

transformed parasitic nematodes meant that RNAi has to be studied by soaking/feeding

nematodes dsRNA. Being obligate parasites, PPNs do not normally feed outside their host

plants. Artificial feeding methods were therefore developed and improved with time to facilitate

ingestion of long dsRNAs or siRNAs by PPNs with the aid of neurostimulants such as

octopamine, resorcinol and serotonin which promote pharyngeal pumping and contractions

(Urwin et al. 2002; Rosso et al. 2005). There has since been limited evidence of successful

RNAi achieved by soaking without the aid of neurostimulants (Kimber et al. 2007; Park et al.

2008). Since J2 nematodes of sedentary endoparasites such as RKNs are the only potential

feeding life stage outside host plant tissues, J2s are the only viable stage to undertake

experiments using RNAi technology by inducing dsRNA ingestion. In vitro ‘soaking’ is

currently the most effective strategy to screen the potential function of genes in a manner that

bypasses the laborious and time-consuming process of plant transformation. It also provides a

quick assay to screen candidate genes for nematode control through HIGS. Soaking J2s in

dsRNA is therefore the most widely used technique for preliminary assessment of gene

knockdown in PPNs.

The function of several effectors and essential genes of sedentary and migratory groups of PPNs

have been evaluated using RNAi (reviewed in Lilley et al. 2012). In Meloidogyne spp., the

focus has been on the genes that are involved in parasitism or nematode viability, but not for

genes involved in the RNAi machinery. Therefore, the aim of this study was to assess how

silencing of small RNA pathway genes affects infectivity of the nematodes and their

development. Twenty seven genes from M. incognita identified in Chapter 2 were analysed after

soaking J2s in dsRNA from them, and phenotypes and levels of infection in tomato plants were

assessed 28 days after the soaking treatments.

60


Identified protein domains for the 27 selected genes were analysed by tblastx against the

nematode database to define the predicted exon positions. The results were then used to design

primers spanning the complete protein domains.

3.3.1 Primer design

Primers were designed using IDT primer quest tool to amplify coding regions of functional

protein domains of the selected genes. Two M. incognita genes involved in cuticle collagen

production (rol-6) and body wall muscle troponin C (pat-10) were also included in the

experiments as phenotypic controls. In addition, dsRNA synthesised from a GFP amplicon of

Aequorea victoria was also included in the experiments as a non-target control. Based on the

similarity to C. elegans genes, primer sequences were named after those of C. elegans, except

for the argonaute coding contig CABB01002242 which significantly matched to several C.

elegans argonautes, and was named ‘2242’ based on contig number. Primer sequences and

expected amplicon sizes are listed in Table 3.1.

Table 3.1: Primers used to amplify genes of the small RNA pathways of M. incognita.

Gene Primer 5′-3′ Protein Domain Contig ID Amplicon (bp)

RNAi and miRNA pathway gene primers

Rsd-3-F

Rsd-3-R

ATTTGCCCCTTCATCTTTTCCTC

ACTGAAACTGAACAAAAAGTTCGTG

ENTH-Epsin CABB01006346 314

Xpo-1-F

Xpo-1-R

TAAGATCGTCAAGCAAGAATG

TGAGATCTATCAGATTGTTCT C

XPO1 CABB01004119 323

Xpo-2-F Xpo-2-R

ATCTAGCTGCGCCAATAAC TGATGTTGCTGAATTTAAACC

CAS_CSE CABB01000462 587

Dcr-1-F Dcr-1-R

TTCCTGCAGGCAAAAGATTGTC GGTACTGTGCAAAATTACCATCTG

DSRBF CABB01000157 251

Drh-1-F

Drh-1-R

GTCAAGCAAGTCGCGAAG

CCCTTCTTTGAACTCTAGCA

Helicase C Terminal

domain

CABB01006008 192

Drh-3-F

Drh-3-R

GTGGTGTCATAACCAAAATGTCTG C

TTG TGC ACC AAC TGG AAG TG

DEAD CABB01002056 284

Drsh-1-F

Drsh-1-R

TAGAATTTCTTGGAGATGCTGTTG

ATCACATTGTTCAAGACCAGAATC

Ribonuclease IIIa CABB01000477 280

Pash-1-F

Pash-1-R

TCAGCATAA ACCTACTCGTG

TCGGCTTTG AATCAATTTCAAG

- CABB01004277 539

Tsn-1-F Tsn-1-R

TTCCCTATCTACAGCGTTCGC CCT CAA CAT GAT GCT TCCATATACC

Staphylococcal nuclease 4

CABB01000055 444

Vig-1-F Vig-1-R

TTTCGGTCGCGCCGTTTTG AATCTTCTCATCAGCTCCTTCGCC

HABP4_PAI-RBP1 CABB01000081 205

Smg-2-F

Smg-2-R

ATGCCATAACGATTGTCTACTC

CAAATTCTGGCCTCGGAC

P loop AAA12 CABB01008394 301

61

3.3.2 RNA extraction

RNA was extracted from freshly hatched J2s and eggs using the Trizol method (Tan et al.

2013). Nematodes were macerated in a 1.5 mL microfuge tube by shaking with stainless steel

beads (3.2mm) in a Tissue Lyzer (Qiagen, Australia) at 25 Hertz for 2 minutes. One mL of

diluted (3:1) Trizol LS reagent (Life technologies Corporation) was added to the extracted total

nucleic acids followed by chloroform purification with 200 µL of chloroform, then centrifuged

at 12,000 g for 10 minutes to separate phases. Clear supernatant was transferred to a clean 1.5

mL microfuge tube and nucleic acids were precipitated overnight by adding 1/20 volume 5M

NaCl and 2.5 volumes of 100% ethanol, followed by centrifugation at 12,000 g for 30 minutes.

The pellet was then washed with 70% ethanol, resuspended in 50 µL of RNase-free water and

subjected to 6.8 Kunits of DNaseI in solution with 10 µL of RDD buffer (RNase-Free DNase

set, Qiagen, Australia). The reaction was incubated at room temperature for 10 minutes and the

nucleic acids purified by chloroform extraction and ethanol precipitation. Resuspended RNA

Gene Primer 5′-3′ Protein Domain Contig ID Amplicon (bp)

Smg-6-F

Smg-6-R

GTATCAATTTATGTAGACGC

TGGAAATGTTCGGACAGG

PIN CABB01000011 131

Ego-1-F

Ego-1-R

CGAACTCAAGAACCTTTTTTCCG

CTGCTCGTTGATGTTTAAGTGC

RdRp CABB01000449 342

Rrf-1-F

Rrf-1-R

TATGCTGACACGCTCCTTAATTG

TCTGCCATGTAATCAAATGCATCC

RdRp CABB01000474 682

Eri-1-F Eri-1-R

GTGATTGATTTTGAATGTAGCTGTG AAGCATCATCCATTCCACAATGTTC

DEDDh 3’-5’ exonuclease domain

CABB01001883 473

Gfl-1-F

Gfl-1-R

AACAGGTTTCTCGTTGACGTCAG

TTCCTCAGAAAAACAGCAAAGGG

Yeats CABB01000795 281

Xrn-2-F Xrn-2-R

CTGTATCTCGATATGAACGG CAAGCATAATCAAGTCTGC

XRN 5’-3’ exonuclease

CABB01001503 553

Rha-1-F

Rha-1-R

TGGGTTTAAGAGGAATTTCTC

CATAAACAACATCATCAATTG

Helicase C Terminal

domain

CABB01000079 669

Mes-2-F

Mes-2-R

CCACATTTTATTGACTGTTGG

TGCGAATGAGGATATCGAGA

SET CABB01002321 258

Mes-6-F

Mes-6-R

TACAGTTGAAAAACTCATACCCC

AACTTTCAGGCCACACTC

WD40 CABB010000967 171

Mut-2-F

Mut-2-R

AACTTACAGGCACTATAATAAC

TGC TGT TGT TGT TCT TTC TTC

PAP-TUTASE CABB01003815 204

Mut-7-F

Mut-7-R

TCCAACACCAATTTT CTCAGCC

TCTGAGCAAGGCCTTTCC

Exoribonuclease

domain

CABB01000055 187

Ekl-4-F

Ekl-4-R

TCGTTGGCCTGAATATAGAC

GATTTGACAATTGGTCAAGC

DMAP1 CABB01002952 245

Alg-1-F

Alg-1-R

CGGATCGAATGACATGTC

ATTCCTGGTTGGTATCCTC

PIWI CABB01000336

476

Csr-1-F

Csr-1-R

CTGAAGTTCATCTTGAGTCA

TTGGACTCAACTACGTTC

PIWI CABB01000355 674

Ppw-2-F

Ppw-2-R

AACCGAAGTCGTCACACA

CTTTGCCGAAATTCCATGTTC

PIWI CABB01001343 661

2242-F

2242-R

GCAGCATAGTGATGTCTAG

GCCAACGCTCTTTA AGG

PIWI CABB01002242

696

Other Primers

Rol-6-F Rol-6-R

GGCTATTGCTTTTAGCGGAGC TGCCATGATCTCCCGACT TCC

Cuticle collagen-Collagen triple helix

CABB01000004 609

Pat-10-F

Pat-10-R

TTCAATCAGTCTCTCCAGCC

AATTCGACGCAGACGGCAG

Ef-hand calcium

binding motif

CABB01000228 303

hpGFP-F hpGFP-R

TAACTCGAGTCTAGATTCACTGGAGTT TACGGTACCGGATCCTAATGATCAGC

- - 524

62

was quantified (Nanodrop ND-1000 Spectrophotometer) and 500 ng used to synthesise cDNA

in a standard 20 µL reverse transcription reaction using the High capacity cDNA synthesis kit

(Applied Biosystems) following the manufacturer’s protocol without an RNase Inhibitor.

3.3.3 Amplification of target genes

Approximately 200 ng of cDNA was used for polymerase chain reactions (PCRs) in a standard

20 µL reaction with 30 cycles in a thermal cycler (Applied Biosystems 2720). The reactions

were made of 5x PCR buffer (final concentration 3mM MgCl2 and 1mM dNTPs) and 0.5 units

of Taq polymerase enzyme (Bioline). Ten picomols each of the forward and reverse primers

were added into the reaction. Cycling conditions were 95 °C for 5 minutes followed by 30

cycles of 95 °C for 30 secs; 55 °C for 30 secs; 72 °C for 30 secs and a final extension step of 7

minutes at 72 °C. To visualise the amplified product, reactions were run on a 1% agarose gel in

1x TAE alongside a 100 bp DNA marker (New England BioLabs).

3.3.4 Cloning of amplicons into RNAi vector

Although multiple domains were amplified for each gene, one domain each for 27 genes and the

three controls (rol-6, pat-10 and gfp) were cloned into pDoubler transcription vector using the

restriction sites AfeI/XhoI and KpnI. The multiple cloning site of pDoubler (Figure 3.1) is

flanked by two T7 promoter sequences and allows a cloned gene fragment to be digested out

with the T7 sequences for in vitro transcription using the enzymes, EcoR1 or NotI. All the

amplicons were first amplified with primers with the restriction sites XhoI and KpnI except for

xpo-1 and xrn-2 where AfeI was used instead of XhoI (Figure 3.1).

Following PCR amplification, the vector and amplicons were digested with 1 U of each enzyme

(New England BioLabs) per µg of DNA at 37 °C for 3-5 hours in a thermal cycler in a double

digest reaction. Digested gene fragments and linearised pDoubler were run on a 1% agarose gel,

cleaned up using the Wizard® SV Gel and PCR Clean-Up system (Promega, Australia)

according to manufacturer’s protocol and quantified using the Nanodrop spectrophotometer.

Standard 10 µL ligation reactions were set up using 50 ng of vector and a 1:3 vector to insert

ratio with 3 U of T4 DNA Ligase (New England BioLabs) and incubated at room temperature

overnight.

63

pDoubler

2617 bp

pUC Origin

HsLev-11 gene

M13-F (7 nt)

M13-R (934 nt)

Kanamycin gene

T7 promoter region

T7 promoter region

BamHI (189)

AfeI (7 7 5)

AscI (182)

AvrII (7 85)

KpnI (199)

XbaI (7 7 9)

Xho I (7 67 )

EcoRI (146)

EcoRI (827 )

NotI (137 )

NotI (834)

Figure 3.1: RNAi cloning vector pDoubler with T7 transcription sites on both sides of the

multiple cloning site (MCS). Template for dsRNA synthesis can be digested out using

restriction sites EcoR1 and Not1. An H. schachti lev11 gene is cloned in the vector to

demonstrate the map of a typical construct.

Escherichia coli JM109 cells (Promega, Australia) were used for all transformations. Cells were

made competent using the rubidium chloride method (Promega Subcloning Notebook, 2004).

Five µL of the ligation reaction for each gene was used to transform 25 µL of E. coli competent

cells using the heat shock method (Promega Technical manual No. 042). Briefly, bacteria and

ligation mix were incubated for 30 minutes on ice followed by 40 seconds at 42 °C in a water

bath. After another two minutes on ice, cells were incubated for 1.5 hours in 700 µL of LB

(Luria-Bertani) broth at 37 °C with vigorous shaking at 225 rpm. The bacterial suspension (150-

200 µL) was then plated on LB agar plates supplemented with 50 µg/mL of kanamycin

monosulfate and incubated at 37 °C for 16-18 hours. Bacterial colonies were screened by PCR

where 5 µL aliquot of a colony resuspended in 20 µL of PCR-grade water was used as DNA

template with 10 picomols of each primer M13-F (5′-TAAAACGACGGCCAGT-3′) and M13-

R (5′-CAGGAAACAGCTATGAC-3′) in a standard 20 µL reaction. PCR conditions and

analysis of amplicons were as described under Section 3.3.3 except that the number of cycles

was 25 and extension time for each cycle was one minute. The remaining 15 µL of bacterial

suspension from PCR positive colonies were grown in 5 mL LB broth supplemented with 50

µg/mL kanamycin monosulfate at 37 °C (16-18 hours) with shaking at 225 rpm. Cloned

plasmids (pDoubler with gene inserts) were recovered from 4 mL of this culture using the

Wizard® Plus SV Minipreps DNA purification system (Promega, Australia) according to

manufacturer’s protocol.

64

3.3.5 Confirmation of sequences cloned into vectors

Purified plasmid DNA was analysed by Sanger sequencing using the facilities at the Western

Australian State Agricultural Biotechnology Centre (Applied Biosystems Industries; ABI 3730

96 capillary machine). ABI BigDye terminator version 3.1 reagents was used for all reactions.

One sixteenth dilution of a full reaction was done using 3.2 picomols of M13-F or M13-R in

separate reactions with 1.75 µL of sequencing buffer, 0.5 µL of dye terminator mix and 300-500

ng of plasmid in a total volume of 10 µL. Reactions were subjected to thermal cycling at 96 °C

for 2 mins, 25 cycles of (96 °C for 10 secs, 55 °C for 5 secs; 60 °C for 4 mins) ending with a

hold at 14 °C. DNA, post-reaction, was transferred to a 600 µL microfuge tube and precipitated

by adding 1 µL of 125 mM EDTA, 1 µL of 3M sodium acetate and 30 µL of 100% ethanol.

After a brief vortex, reactions were spun down and, incubated at room temperature for 20

minutes in the dark followed by centrifugation at 13,000 rpm for 30 minutes. Supernatant was

removed and replaced with 125 µL of 70% ethanol to wash the pellet. After centrifugation for

another 5 minutes, the supernatant was discarded and pellet dried in the dark for 20 minutes

before submitting for sequencing. Sequencing results were analysed using FinchTV 1.4.0

(Perkin Elmer, Inc.). A nucleotide blast (blastn) was done in NCBI (http://www.ncbi.

nlm.nih.gov/) against whole genome sequence database of M. incognita for sequence

verification.

3.3.6 Synthesis of dsRNA

After sequence confirmation, cloned fragments with T7 sequences at the 5′ and 3′ ends were

digested out of pDoubler using EcoRI for all the sequences of the genes except dcr-1 and drsh-1

for which NotI was used as the sequences had EcoR1 sites. HiScribe T7 In vitro transcription kit

(New England BioLabs) was used to generate dsRNA from 1 µg of template for each gene in a

standard 20 µL reaction. In vitro transcription reactions were treated with DNaseI to remove

template DNA. DsRNA was purified using chloroform extraction followed by NaCl and ethanol

precipitation as described in Section 3.3.2. DNA-free dsRNA was pelleted and resuspended in

30 µL RNase-free water. After quantification (Nanodrop) it was ran on 1% agarose gel in 1x

TBE to assess the quality. The dsRNA of the respective genes was named as ‘dsgene’ for this

study.

3.3.7 In vitro feeding of dsRNA to nematodes

Freshly hatched J2s were used for all soaking experiments. Egg masses were collected from

roots of infected tomato (cv. Grosse Lisse) in water in a 1.5 mL microfuge tube. These were

washed with 2% sodium hypochlorite for 2 minutes and subsequently washed four times with

water to remove the gelatinous matrix and residual sodium hypochlorite by pelleting eggs at

65

12,000 g for 1 minute. The eggs were transferred into 5 mL tubes with 2 mL of water and

incubated at 25 °C with gentle agitation (60 rpm) for two days. Hatched J2s were collected after

passing through a 38 µm sieve which allows active M. incognita J2s to migrate through, leaving

dead nematodes and other debris. The collected nematodes were used for dsRNA soaking

experiments in 70 µL M9 buffer (3 g KH2PO4, 6 g Na2HPO4, 5 g NaCl, 1 mL 1 M MgSO4)

containing 1 mg/mL of dsRNA supplemented with 50 mM octopamine (octopamine

hydrochloride), 3 mM spermidine (spermidine trihydrochloride) and 0.05% gelatine. All stock

solutions were made with sterile M9 buffer. Tubes containing the soaking solution with 7,000

J2s each were incubated at 25° C for 16 hours in the dark. Controls contained 1 mg/mL of dsgfp

or no dsRNA. FITC was used at a concentration of 1 mg/mL in parallel to monitor solution

uptake, supplemented with the same soaking solution without dsRNA. After 16 hours

incubation, nematodes were removed from the soaking solution washed before further

assessment and 20 µL of the soaking solution was run on a 1% agarose gel to assess dsRNA

integrity.

3.3.8 Observation of RNAi phenotypes

After 16 hours soaking in dsRNA, any effects on nematode behaviour were observed and

assessed using an Olympus BX-51 microscope. Nematodes were viewed in bright field while

FITC uptake was monitored using the FITC filter (450-480 nm). One thousand nematodes were

observed for each reaction at 4x and 10x magnifications. The control nematodes used to

compare phenotypic effects of soaking were fed no dsRNA and dsgfp. Because there was no

previous reports of RNAi phenotypes for any of the small RNA pathway genes in parasitic

nematodes, pat-10 and rol-6 were used as ‘positive’ controls. These genes are required for

cuticle and muscle development and down-regulation results in a physically observable

phenotype (Adamo et al. 2012; Nsengimana et al. 2013)

3.3.9 Assessment of nematode infectivity and development

After soaking for 16 hours, the ability of nematodes to infect plants was assessed. Tomato seeds

(cv. Grosse Lisse) were germinated on moist Whatman filter paper in 15 cm petri plates.

Germinated seedlings were transferred into white sand in 5 x 4 trays each with a capacity of 120

cm3. Ten 2 week old seedlings were infected with 400 J2s each from the dsRNA-fed and control

(dsRNA to gfp and no dsRNA) treatments. Plants were watered sparingly the first two days after

infection. Plants were grown under glasshouse conditions (25 ± 5 °C) for four weeks and then

gently uprooted and washed thoroughly with tap water to count galls formed on the roots. The

number of galls present on 6-7 plants per treatment was counted using a dissecting microscope.

The remaining 3-4 plants for each treatment were allowed to grow for another three weeks after

66

which females were extracted using fine forceps to observe morphology. After counting the

galls, roots were separated from the shoots at the collar and dried at 50-55 °C for 18 hours in a

drying oven. Dried roots were weighed and used to calculate infection severity expressed as

number of galls/gram of dry root weight. Morphological differences were observed using a

dissecting microscope and compared to those developing on plants infected with control

nematodes soaked with dsgfp or no dsRNA.

3.3.10 Gene expression of target genes

Gene knockdown was assessed on nematodes 16 hours after feeding with dsRNA. Two

thousand J2s from each feeding experiment were washed thoroughly with DEPC-treated water,

snap frozen in liquid nitrogen and stored at -80 °C to prevent RNA degradation until further

processing. RNA for these nematodes was extracted with the Trizol method followed by DNase

treatment as described in Section 3.3.2. cDNA was synthesised with 100 ng of RNA for each

treatment and amplified with gene specific primers at 30 cycles with 1 in 10 dilution of cDNA

in a 10 µL PCR reaction for a preliminary analysis of transcript produced after 16 hours of

dsRNA feeding. Reactions were run on a 2% agarose gel alongside a 100 bp DNA marker for

comparative analysis of transcript abundance. M. incognita 18S ribosomal RNA (rRNA)

primers (18S-F 5′-TAGAGGGATTTGCGGCGTTC-3′, 18S-R 5′-GGTTTACCCGCCCCTTT

CAG-3′) were used to amplify 18S transcript as normalisation control. Relative band intensity

was analysed visually in control and dsRNA-fed nematodes.

3.3.11 Statistical analysis

SPSSv20 software package (IBM Corporation, US) was used for analysis of variance (ANOVA)

and calculation of means, standard deviation and standard error. Significance between

treatments was tested at p<0.05 and pair-wise comparisons were done post-hoc using the

Tukey’s test. Microsoft Excel Analysis ToolPak was used for construction of bar charts with

error bars representing standard error for each treatment.

3.4 Results

3.4.1 Phenotypic effects of in vitro RNAi of target genes

Amplified products were sequenced and compared to the exon regions of the genomic contigs

that were used to design primers. Blastn analysis showed 100% similarity of each of the

amplified fragments to contigs of M. incognita and between 87-95% to those of M. hapla, with

perfectly complementing fragments (>25 bp) within the amplicons. When the feeding solution

was run on a 1% agarose gel there was no indication of dsRNA degradation after the 16 hour

treatments.

67

FITC uptake was observed 16 hours after feeding J2s. When eggs were present, FITC was

detected in most, suggesting that at least the outer layers were permeable to the feeding solution

(Figure 3.2). A small percentage (<10%) of nematodes soaked with either no dsRNA or dsRNA

to gfp were straight and showed little movement. In contrast, RNAi of rol-6 resulted in vigorous

movement of treated nematodes which appeared to be involuntary. Those fed with dsRNA to

pat-10 were curled and appeared paralysed.

After soaking in dsRNA, the observed phenotypes for the 27 genes studied could be divided

into three major groups: first, those with straight bodies showing little movement, mainly

restricted to the head region, with the rest of the body paralysed. This group included genes fed

with dsRNA targeting rsd-3, xpo-2, dcr-1, drh-1, pash-1, tsn-1, vig-1, ego-1, eri-1, mut-2, alg-1,

ppw-2, csr-1 and 2242. The second group consisted of J2s with wavy bodies and was typical of

J2s ingesting dsRNA of drh-3, drsh-1, rrf-1, smg-2, smg-6, xrn-2, rha-1 and ekl-4. This wavy

form seemed to be a result of some sort of paralysis, except for smg-6 for which the wavy form

was observed with jerky movements in the nematodes’ bodies. The third group consisted of J2s

curled with little movement and this was observed for nematodes fed with dsRNA of mut-7 and

xpo-1.

J2s fed dsRNA complementary to genes dcr-1, drh-1, drh-3, pash-1, vig-1, ego-1 and the

argonaute coding sequence 2242 were generally straight and paralysed, except for drh-3 which

displayed the wavy paralysed phenotype but also resulted in dead nematodes. The percentage of

dead nematodes ranged from 10% to 60% with ~60% resulting from feeding with dsRNA of

drh-1, pash-1 and 2242 while about 50% J2s fed dsego-1 and dsvig-1 were dead. Interestingly,

four out of six of these genes encoded helicases. FITC uptake by J2s and prominent phenotypes

after 16 hours of dsRNA ingestion are presented in Figure 3.2.

Seven functional groups i.e. transport proteins, dicer complexes, RISC proteins, amplification

proteins, RNAi inhibitors, nuclear RNAi proteins and argonautes were targeted by RNAi in this

experiment. The RNAi phenotypes observed were not typical of genes from any particular

group except RNAi of the four argonautes which resulted in nematodes with straight body

phenotype with paralysis 16 hours after feeding. All soaking treatments affected nematodes

phenotypically, except for nematodes treated with dsmes-2 and dsmes-6, which appeared

similar to controls. All phenotypes observed after 16 hours of feeding dsRNA complementary to

the target genes are presented in Table 3.2.

68

500 µm

No dsRNA dsxrn-2

dsdcr-1 dsego-1

dsvig-1

dsmut-7

500 µm

dstsn-1

ds2242 dsrol-6

500 µm

dsgfp

FITC

500 µm

FITC

dsalg-1

dsdrsh-1

FITC

200 µm

Figure 3.2: RNAi phenotypes of M. incognita J2s observed after 16 hours of feeding

with dsRNA of selected genes. FITC uptake was also recorded (Labelled FITC) in J2s.

Eggs in the solution also absorbed some FITC. Phenotypes presented after feeding

dsRNA to different target genes (ds‘gene’ refers to dsRNA to the target gene i.e. no

dsRNA, dsgfp, dsxrn-2, dsego-1, dsdcr-1, dsmut-7, dsvig-1, dsalg-1, dstsn-1, ds2242,

dsdrsh-1 and dsrol-6. Arrows indicate J2s with prominently different phenotypes from

the controls (controls were: no dsRNA and dsgfp).

69

Table 3.2: Phenotypes observed in J2s of M. incognita after 16 hours of soaking in dsRNA (1

mg/mL) of genes involved in the RNAi pathway.

3.4.2 RNAi of target genes of nematodes reduces host infection

Nematodes soaked in dsRNA for 16 hours were used to infect tomato plants to assess their

infectivity after treatment. Experiments were conducted in batches with respective controls and

statistical significance tested accordingly. There were significant differences in the levels of

infection between nematodes fed with dsRNA of complementary target genes and those soaked

in control solutions without dsRNA or dsRNA to gfp.

After infection with treated nematodes, tomato plants were maintained in a glass house, and

roots were harvested four weeks later. The level of infection was assessed by the number of

galls formed per gram dry root weight, to account for differences in sizes of roots masses: the

Gene J2 Phenotype

rsd-3 Straight body. Paralysed slow movement.

xpo-1 J2s inactive with curled bodies.

xpo-2 Straight body.

dcr-1 20% dead. Straight stiff body. Only head moving.

drh-1 60% dead. Straight paralysed body.

drh-3 20% of J2s dead. Wavy movement

drsh-1 Abnormally wavy rigid body with movement only in the head.

pash-1 60% dead. Straight body and non-motile.

tsn-1 Straight body. Slow movement.

vig-1 50% of J2s dead. Straight body.

rrf-1 J2s alive with slow movement. Abnormally wavy bodies.

ego-1 50% dead. Straight appearance with very slow movement.

smg-2 Wavy but stiff bodies.

smg-6 Wavy body. Faster jerky movement.

eri-1 Straight body, slow movement.

gfl-1 J2s alive with slow movement.

mut-7 Slow movement with curling bodies.

xrn-2 More wavy bodies than normal. Very slow movement.

mut-2 Straight body. Mostly alive but with slow movement.

mes-2 J2s active and similar to controls.

mes-6 J2s active and similar to controls.

rha-1 Vigorous movement. Wavy body.

ekl-4 Wavy and slow movement.

alg-1 Straight body.

ppw-2 Straight body. Slow movement.

csr-1 Straight body. Some show body curling with jerky movement

2242 60% dead. Paralysed with straight body.

rol-6 Vigorous movement. Moving faster than controls

pat-10 Wavy, slow movement. Curling body.

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results obtained for the different treatments are presented in Figure 3.3. The lowest number of

galls observed was by nematodes fed with dsdcr-1, with a reduction of up to 90% compared to

treatments lacking dsRNA. Treatments thought to disrupt expression of the miRNA processor

complex (drsh-1, pash-1) and the RNAi inhibitor (gfl-1) resulted in ≥ 70% reduction in

infection. For the genes xpo-1, xpo-2, drh-1, vig-1 ego-1, mut-7, rha-1, 2242 and pat-10, RNAi

reduced nematode infectivity by ≥ 50%. Targets with less RNAi effect on nematodes were drh-

3, xrn-2, mut-2, mes-2, ekl-4, the three argonautes (alg-1, ppw-2, csr-1) and rol-6, with a ≥ 25%

reduction in infection. The fourth argonaute (2242) however, reduced nematode infestation by

68%. The three helicases of the dicer complex reduced infection by between 45-90%.

RNAi of the two major transport proteins for miRNA pathway xpo-1 and xpo-2 affected

nematode infectivity significantly (61% and 53% reduction respectively) unlike rsd-3 where

there was only 24% reduction in infectivity. For the group of nuclear RNAi effectors, disruption

of rha-1 function resulted in reduction in plant infection by 64%, but there was no change in

infectivity for dsmes-6 soaked nematodes. Notably, apart from ego-1, RNAi of amplification

genes (smg2, smg-6) did not seem to have significant (p<0.05) effect on nematode infectivity.

The other RdRp rrf-1 when disrupted resulted in a significant (p<0.05) increase in infection

(19%): the only gene for which this pattern (increased infection) was observed.

One important observation was that egg masses were produced earlier as compared to controls

(no dsRNA and dsgfp) for nematodes treated with dsgfl-1, dseri-1 and dsrrf-1. For these, egg

masses were observed four weeks after infection whilst controls had no egg masses (Figure

3.4A). For the feeding experiment for ego-1 and mut-7, fungal infection was observed for the

plants and the galls produced by nematodes were deformed as indicated in the Figure 3.4B. Egg

masses produced on the roots lacked the characteristic swollen gall-like appearance and number

of galls reduced by 73% for ego-1 and 20% for mut-7 (data not shown). The experiment was

repeated to further investigate the deformed galls and this time normal galls were produced for

that experiment with reduction in infection of 62% for dsego-1 and 55% for dsmut-7 (Figure

3.3).

No such deformity was observed for dsRNA treatment of the other genes used in this study.

Interestingly, for most of the genes where RNAi did not reduce infectivity of nematodes (tsn-1,

rrf-1, xpo-2, eri-1, smg-2 and smg-6), there were nevertheless phenotypic changes in the

appearance and movement of nematode 16 hours after feeding.

71

Figure 3.3: Average number of galls per gram of dry root of plants infected with dsRNA-fed nematodes. Genes belonging to seven

functional groups were targeted together with controls. (Different colours of bars represent different groups of genes of the small RNA

pathway of nematodes as indicated in the figure legend). Significance is represented by * at p<0.05 with respect to the no dsRNA control

for the separate experiments. Bars represent mean (n= 6-7) ± standard error.

72

(A)

(B)

3.4.3 RNAi effects on nematode development

dsmut-7 dsego-1

dseri-1

dsgfl-1 dsgfp No dsRNA

dsrrf-1

Figure 3.4: (A) Infection on tomato roots by M. incognita J2s after soaking without dsRNA, or

with dsgfp, dsgfl-1, dsego-1 and dsrrf-1. Egg masses are indicated by arrows on the surface of

galls produced by nematodes treated with dsgfl-1, dsego-1 and dsrrf-1 but are absent from

controls (no dsRNA and dsgfp). Eggs were stained with phloxine-B. (B) Deformed/no galls

produced by nematodes fed on dsego-1 and dsmut-7. Arrows indicate egg masses on root

surface.

73

Adult females developing in tomato roots were dissected out of galls and compared with those

of controls. Morphological differences were observed in females soaked in solutions containing

dsRNAs at the J2 stage. Females which developed from J2s previously soaked in dsdrh-3 and

dsmut-7 had elongated bodies (fusiform) instead of the wild-type spherical saccate posterior.

This kind of deformity was observed for 41% and 42% females from treatment with dsdrh-3 and

dsmut-7 respectively. For the nematodes soaked in dsvig-1 and dsmut-2, 30% and 85% of

developed females respectively were smaller than those soaked with dsgfp or no dsRNA. In

addition, developed females of J2s previously fed with dsRNA corresponding to genes with

roles in the miRNA pathway e.g. xrn-2, xpo-2, pash-1, drsh-1, and alg-1 were mostly abnormal.

In particular, RNAi of xpo-2, mut-2, rha-1 and 2242 resulted in females that were translucent

compared to those of controls that appeared pearly white (Figure 3.5). However, there was

variable number of nematodes depicting this deformity i.e. it was 100% for dsmut-2, 50% for

dsxpo-2, 75% for dsrha-1 and 30% for ds2242.

Interestingly, nematodes fed with dssmg-2 showed a wavy paralysed phenotype after soaking in

dsRNA, but no significant reduction (p<0.05) in infectivity was found. However, their

development seemed to be affected since abnormal females were present after four weeks of

infection on tomato plants (Figure 3.5). Females developing from J2s soaked in dsmut-2 were

not only deformed, but were also translucent: the J2s appeared affected after feeding and there

was a 40% reduction in infection. Severely paralysed nematodes as a result of soaking in dstsn-

1 and those resulting in abnormally wavy bodies in response to dssmg-2 and dssmg-6 showed

no reduction in plant infestation. Similarly soaking in dsRNA to smg-2, alg-1, ppw-2, csr-1 did

not result in greatest reduction in infection but abnormal female development was observed. No

relationship could be established between phenotype of J2s after dsRNA feeding, plant infection

and abnormal female development.

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dsalg-1

2.0mm

ds2242

2.0mm

dsxpo-2

2.0mm

dsmut-2

2.0mm

dsvig-1

2.0mm

dspash-1

2.0mm

dsdrh-3

dssmg-2

2.0mm

dsrha-1

2.0mm

2.0mm

dsxrn-2

2.0mm

dsgfp

2.0mm

No dsRNA

2.0mm

Figure 3.5: M. incognita females dissected from tomato roots following infection with J2s

treated without dsRNA, and with dsgfp, dsxrn-2, dsxpo-2, dsdrh-3, dsmut-2, dsvig-1,

dssmg-2, dspash-1, dsrha-1, dsalg-1 and ds2242, seven weeks after plant infection. Arrows

point to abnormal females in different treatments compared to controls. Scale bar = 2 mm.

75

3.4.4 Transcript abundance after in vitro feeding

PCR was used to assess transcript abundance in nematodes fed by dsRNA. For this, the

nematodes from dsRNA soaking treatments which showed a reduction in infection or changes in

the morphology of adult females were further analysed. Amplification (PCR band intensity) of

target genes from cDNA made from similar amounts of RNA derived from nematodes treated

with dsRNA and no dsRNA or dsgfp were compared. Amplification of 18S rRNA transcript

was used as internal control.

It appeared that 18S rRNA gene transcription was affected by soaking in dsRNA (both up-

regulation and down-regulation was observed for dsgene and dsgfp treated nematodes) but this

was not consistent for all experiments (Figure 3.6). Furthermore, all of the genes tested seemed

to be up-regulated by dsRNA feeding, except drsh-1 and alg-1 which displayed no apparent

change in transcript abundance. The transcripts up-regulated were of the genes gfl-1, vig-1,

pash-1, xpo-2, 2242, xrn-2, mut-2, mut-7 and pat-10 with the greatest up-regulation for dsgfl-1

and dsxrn-2. To test whether this amplification was a result of dsRNA that was fed to the

nematodes and remained bound to their body and extracted during RNA extraction, primers

designed from a region of genes other than the part used to generate dsRNA were also used for

PCRs. The result was not different as there appeared to be a higher intensity of amplicons

compared to those of the controls. Some genes could not be amplified from the cDNAs of J2s

fed with no dsRNA, dsgfp and dsRNA of some genes even when the PCR cycles were increased

to 35: these included cDNAs of J2s fed with dsRNAs of dcr-1, rol-6, rha-1, csr-1 and ppw-2

(Figure 3.6). Expression of 18S rRNA for these was high and appeared as bright bands after 30

cycles. The same primers when tested against cDNA extracted from mixed stage nematodes

amplified the gene products. This implies that the expression of these genes was very low in M.

incognita J2 stage nematodes, and so was not readily detected from the amount of template

cDNA used in the PCRs.

76

Figure 3.6: RT-PCR amplification of target genes in control and dsRNA-fed J2s with respective gene

specific primers and 18S rRNA primers.

dsRNA

treatment

18S rRNA primer

Gene specific primer

No dsRNA dsgfp dsgene No dsRNA dsgfp dsgene

drsh-1

dcr-1

No amplification

gfl-1

vig-1

pash-1

alg-1

xpo-2

2242

xrn-2

mut-2

mut-7

pat-10

rol-6

No amplification

rha-1

No amplification

csr-1

No amplification

ppw-2

No amplification

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3.5 Discussion

The primary aim of this study was to assess infectivity and development of M. incognita J2s

following treatment with dsRNA of genes involved in the small RNA pathway. For the 27 genes

used, the ones for which there were definite phenotypes after soaking were mut-7 (curling

body), rol-6 (vigorous uncontrollable movement), vig-1, tsn-1, ego-1, dcr-1 and the four

argonautes (straight paralysed body). The straight phenotype displayed by the J2s after 16 hours

of soaking in dsRNA for most of the genes might also be related to the stress imparted by gene

knockdown treatment rather than a direct effect of RNAi of the target gene, since some J2s in

control reactions (no dsRNA and dsgfp) also showed this phenotype. It is possible that

continued stimulated feeding and gene knockdown could overwhelm the RNAi machinery, and

this could contribute towards treated nematodes going into a ‘state of rest’, hence inactive and

straight. Similar straight phenotypes have been reported before as a result of long term exposure

(7 days) to dsRNA corresponding to the neurotransmitter FMRFamide-like peptides, but not

when the same treatment was short-term (24-48 hour) for Globodera pallida (Kimber et al.

2007). Genes involved in the RNAi pathway of nematodes are also related to developmental

regulation (Zhuang and Hunter 2012). A disruption of these mechanisms possibly induces stress

apart from that imparted by dsRNA soaking, which is phenotypically expressed as

inactivity/paralysis.

Several published reports suggest that in vitro dsRNA feeding can be transient. Recovery of

nematodes within 24 hours after washing to remove the soaking solution has been reported for

other nematode species as well, such as Meloidogyne graminicola (Nsengimana et al. 2013),

and after 24 hours soaking for Pratylenchus coffeae, in which the recovery rate for pat-10

transcript was double that of unc-87 after 24 hours soaking with dsRNA (Joseph et al. 2012).

For M. incognita, down-regulated calreticulin and polygalacturonase genes recovered at

different rates after RNAi treatments (Rosso et al. 2005). In this study, in agreement with these

observations, seemingly inactive nematodes were subsequently able to infect plants, with no

difference in gall formation compared to controls – one such example is the gene tsn-1. This

implicates recovery of the J2s following in vitro treatment, and that they were not dead even

though they were inactive after soaking.

Plant infection after soaking for 16 hours was variable: it depended very much on the gene that

was being silenced. The possible reason why the greatest reduction in infectivity after soaking

with dsdcr-1 and dsdrsh-1 was observed may be because dcr-1 is the only dicer-like gene in

nematodes, and it is responsible for all small RNA processing, and drsh-1 is the microprocessor

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for miRNAs before they exit the nucleus so any disruption may affect the overall health of the

organism because of disrupted gene regulation. The infectivity of nematodes after different

treatments could also be affected by factors such as the recovery rate of transcript for the

particular gene, possible presence of multigene families and actual importance of the tested gene

in nematode metabolism. Nevertheless, the results in general show pronounced reduction in

infectivity after soaking, and developmental abnormalities in adult females. Since the greatest

effects were found for genes associated with the miRNA pathway, it may be that after treatment,

longer times were required for the nematode to completely recover. In case of Heterodera

glycines, J2s took 5 to 15 days to recover post dsRNA treatment (Bakhetia et al. 2007).

The formation of giant cells and galls in host plants follows a defined and specific pathway

(Jones and Goto 2011). If the metabolism of J2 nematodes is disrupted as a result of down-

regulation of a gene in a vital pathway, then it is perhaps not surprising that such disturbances

can be manifested in abnormalities in the host response to nematode feeding, resulting in both

abnormal development of nematodes, and abnormal plant responses, manifested by changes in

gall formation, and probably giant cell size and function. The deformed galls observed after

RNAi treatment for ego-1 and mut-7 may therefore be a consequence of silencing of vital

nematode genes in its ability to interact with plant cells, so affecting gall development. The fact

that when this particular experiment was repeated the same result was not obtained may be a

result of seasonal variability of plants or experimental variation of in vitro RNAi, as has been

previously reported for the animal parasitic nematode Ostertagia ostertagi (Visser et al. 2006),

and seasonal differences can affect nematode metabolism, as found for the migratory plant

parasite Pratylenchus vulnus (Britton 2001). Another factor was that in the case where normal

galls were not formed after RNAi treatment there was some fungal infection. This may be co-

incidental or causal.

Adult females that exhibited abnormal development were found after RNAi treatment of J2s for

10 of the 27 genes tested here. Since the miRNA pathway is important in controlling gene

expression, disturbing the balance of the miRNA pathway genes can have an array of knock-on

effects on other gene products (i.e. enzymes) involved in many metabolic pathways of treated

nematodes, such as those involved in moulting, reproduction etc, and such effects might also

explain the early egg mass production after some treatments. Formation of a translucent body is

a strong indicator of affected development and/or poor nutrition of treated nematodes, and

would be accompanied by reduced viability and reproduction fitness. Failure to develop the

typical saccate body shape by M. incognita adult females after in vitro RNAi treatments of J2s

is not new, for example this phenomenon was found for dsRNA treatment targeting a gene

encoding cathepsin L cysteine proteinase in M. incognita (Shingles et al. 2007). However, the

79

translucency of adult females as described here has only been reported in nematodes feeding on

plants producing dsRNA complementary to nematode splicing factor and integrase genes

(Yadav et al. 2006). There may be a pattern in terms of pathways, since the genes targeted by

Yadav et al. (2006) disrupted similar essential cellular processes involved in gene regulation,

and here the translucent appearance of adult females was largely found after RNAi of genes

involved in the miRNA pathway (xpo-2, pash-1, alg-1, xrn-2).

A feature that deserves more study is that, after soaking in dsRNA of target genes followed by

washing and transfer to wild-type tomato plants, a reduction of infection and abnormal

phenotypes of adult females was found. However, despite the abnormal phenotypes, after

soaking in dsRNA, there was not a reduction in transcript abundance, in fact transcript

abundance increased. This phenomenon, that is, an initial increase in transcript level, has also

been found in some instances after RNAi soaking treatments, and it appears to reflect a

feedback mechanism similar to that in the miRNA pathway (Bakhetia et al. 2008). Since the

miRNA pathway is an important component of the regulation of gene expression, some possible

explanations for this phenomenon can be suggested. It may be that in response to perturbations

caused by RNAi treatment, there is a rapid response to increase transcription of targeted genes.

Alternatively, a higher concentration of dsRNA may be required to reduce transcript levels. The

concentration of dsRNA/siRNA required to knockdown a gene probably depends on a number

of factors, such as the specific target and sequence chosen, whether it is part of a multi-gene

family, and whether there are alternative compensatory pathways, as well as differences

between different nematode species (reviewed in Lilley et al. 2012). Regardless of the reports in

which in vitro RNAi effects were temporary (i.e. there was a rapid recovery after treatment), in

other research, concentrations of as low as 0.1 mg/mL have been reported as effective, for

example in knockdown of FMRFamide-like peptides (FLPs) in Globodera pallida, and the

effect increased with increase in RNA concentration (Kimber et al. 2007). The most probable

explanation for an increase in the target mRNA after soaking in dsRNA to that target, is that the

dsRNA concentration (1 mg/mL) used in the experiments initiated a feedback mechanism for

gene regulation which resulted first in a reduction in the presence of target gene mRNA,

followed by a stimulation of transcription of the target gene, but during the recovery phase and

subsequent plant infection, the secondary siRNAs produced by the nematode itself through the

RNAi amplification system then reduced the transcript levels, and so affected and disturbed

growth and development. For other target genes, there are many reports of a reduction of

transcript level long after soaking treatment. For example, in M. incognita the greatest reduction

in target message was observed 20 hours after a four hour dsRNA treatment for the gene Mi-crt,

and the greatest silencing was found 44 hours after soaking for the gene Mi-pg-1 (Rosso et al.

2005).

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The 18S rRNA gene is often used as a consistent reference gene or ‘housekeeping gene’ for

normalisation of comparative transcript analysis in many systems (e.g. Goidin et al. 2001; Bas

et al. 2004). Although its expression may be stable in other animal systems, this gene shows

variable expression levels under certain experimental conditions (Nelissen et al. 2010; De Santis

et al. 2011). In our experiments, inconsistencies in the expression of 18S rRNA genes may be

attributed to the different proportions of live nematodes in different treatments that were used

for RNA extraction, or a response to imbalance in gene regulation caused by dsRNA treatment

targeting RNAi pathway genes themselves.

In conclusion, in vitro RNAi of genes involved in the small RNA pathway of M. incognita J2s,

after soaking treatment and transfer to tomato plants, resulted in reduced viability and

infectivity, and deformities in adult females that developed. Some treated M. incognita J2s were

able to recover from the stress of soaking in dsRNA and were able to infect and feed in host

plants. RNAi of genes involved in the miRNA pathway (drsh-1, pash-1, xpo-2, alg-1, xrn-2)

and helicases of the Dicer complex (dcr-1, drh-1, drh-3) seemed to have comparatively higher

impact on disrupting nematode infectivity and development, hence these genes may be the best

targets to progress for study of plant-delivered dsRNA. The RNAi screens of target genes

undertaken here show that if target genes are to be pre-screened by soaking in homologous

dsRNA, optimisation of conditions is required for factors such as the concentration of the

dsRNA used, the particular target gene and sequence used, whether there is a ‘recovery’

phenomenon after treatment. One variable is that for host-delivered dsRNA, the concentration

present in cells is not known, or the effect of long term delivery of dsRNA. Nevertheless

dsRNA treatments for target genes still provides a valuable filter to decide which targets to

choose to take forward for in planta RNAi studies for nematode control.

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Chapter 4

Host-induced gene silencing of RNAi effectors confers

resistance against Meloidogyne incognita and affects

development

82

Host-induced gene silencing of RNAi effectors confers resistance against

Meloidogyne incognita and affects development




4.1 Abstract

RKNs (Meloidogyne spp.) have broad host range and are the most damaging PPNs. In planta

RNAi provides a potential method of delivery of dsRNA for silencing or reducing specific gene

expression. The transgenic plants generated for these experiments transcribed hairpin (hp) RNA

corresponding to nematode genes from different steps of the RNAi pathway. These were part of

the dicer complex (dcr-1, drh-3), microprocessors (drsh-1, pash-1), nuclear RNAi effectors

(mut-7, rha-1) and the RISC gene vig-1. Expression of hp RNA in these plants reduced

nematode infestation by up to 89% for drsh-1, 76% for mut-7, 74% for drh-3, 71% for dcr-1,

66% for rha-1 and 64% each for pash-1 and vig-1. Host-delivered RNAi also impaired normal

development of nematodes resulting in smaller sized and fusiform females. These results

indicate that longer-term delivery of dsRNA throughout the nematode’s life cycle induces RNAi

of these genes, which significantly affects the nematodes’ ability to enter, migrate in and form

feeding cells in host plant roots, and to develop normally. As a result the target genes studied

could be used as targets for developing resistance in crop plants.

Keywords: Host-induced gene silencing, RNAi pathway, root-knot nematodes, transgenic

plants, nematode control.

4.2 Introduction

Root-knot nematodes establish a sophisticated relationship with cells of host plants in that they

are able to modify the host’s metabolism and establish giant cells at a feeding site. Currently,

there is no one environmentally safe and economically feasible method to control PPNs and

especially RKNs, whose control has been described as a “never ending battle” (Sikora and

Fernandez 2005). HIGS has been studied in a number of plant host-pest/pathogen situations,

including VIGS or incorporation of a transgene producing a hairpin trigger which is processed

into siRNAs by the plant RNAi machinery (reviewed in Dutta et al. 2014).

Arabidopsis thaliana is a good subject for plant transformation studies because of its rapid life

cycle, diploid genetics, its well characterised small genome (125 Mbp) and the fact that it is

83

relatively easy for generating mutants and transgenic plants. Although it is not the best host

plant for cyst and root-knot nematodes, nevertheless the nematodes can complete their life

cycles on Arabidopsis, making it a suitable system for study of the biology of infection of these

nematodes (Sijmons et al. 1991). As a result, Arabidopsis has become a standard host plant to

study host-delivered RNAi to root-knot nematodes: most research using this system has been on

genes involved in nematode parasitism and development.

In Chapter 3, the effects of dsRNA treatment of 27 genes involved in the RNAi and miRNA

pathways of RKNs were assessed using the ‘soaking’ method. From this work, 11 genes that

caused the greatest reduction in infection of tomato plants after dsRNA treatment, and/or

abnormal adult female morphology were chosen for plant-mediated RNAi to study the longer

term effects of continuous ingestion by M. incognita of dsRNA from giant cells.


4.3.1 Cloning of hairpin expression cassettes

The genes chosen for HIGS were dcr-1, drh-3, vig-1, ego-1, mut-7, drsh-1, pash-1, rha-1, alg-1,

2242 and rol-6. The non-target gene gfp and an empty vector with no gene were also used as

controls. The nematode gene sequences and gfp were analysed with the software ‘dsCheck’ to

identify potential off-target effects in the host plant A. thaliana (Naito et al. 2005).

Cloning vector pCleaver provided by Dr. John Fosu-Nyarko was used to prepare the hairpin

expression cassette. pCleaver is a 4 Kbp vector which makes it easier to manipulate through

cloning cycles because of the small size (Figure 4.1). The hairpin cassette in pCleaver has a

constitutively expressing Cauliflower Mosaic Virus (CaMV) 35S promoter and a Nopaline

Synthase (NOS) terminator sequence. The vector confers resistance to the antibiotic kanamycin

in bacteria. Sense and antisense fragments were cloned in tandem, separated by a bean catalase

gene intron (190 bp) between the promoter and terminator sequences, in a two-step process.

Restriction digestion, ligation, transformation, PCR conditions and plasmid purification were

done as described in Section 3.3.4.

Briefly, for each target gene, the 5′− 3′ (sense) fragments were digested out of pDoubler using

the enzymes XhoI and KpnI, except for xpo-1, xrn-2 and gfp for which AfeI was used with KpnI

and ligated to pCleaver linearised by the same restriction enzymes. After ligation, E. coli JM109

was transformed with the DNA using the heat shock method (Section 3.3.4). Colonies were

grown on selection media i.e. LB and 50 µg/mL of kanamycin monosulfate. Colony PCR using

the primers S35S (5′-GATTGATGTGACATCTCCACTGA-3′) and SIntron (5′-TCATCATC

ATCATAGACACACGA-3′) was done to select positively transformed colonies which were

84

then cultured in 5 mL LB broth with 50 µg/mL kanamycin. The cloning vector pCleaver with

restriction enzyme sites and primer positions is presented in Figure 4.1.

Figure 4.1: pCleaver cloning vector. Restriction enzyme sites and primer positions are

indicated. MCS (A) and MCS (B) are multiple cloning sites for ligating target sequences in 5′−

3′ and 3′− 5′ orientations.

Plasmid DNA was purified from cultures and restriction digestion with XhoI and PstI used to

confirm successful cloning. This vector with ligated sense fragment was then linearised using

XbaI and BamHI and the 3′− 5′ antisense fragments of the target genes digested out of pDoubler

with the same restriction enzymes was ligated to the vector. For gfp antisense cloning, the

restriction sites used were AvrII and AscI. After transformation and culture on selective media,

colonies were screened by PCR using primers ASIntron (5′-TCGTGTGTCTATGATGATGA

TGA-3′) and ASNosA (5′-CATCTCATAAATAACGTCATGCATT-3′). PCR positive colonies

were cultured in LB broth with kanamycin (50 µg/mL) overnight and plasmid DNA purified.

The DNA was digested with NotI to confirm successful cloning and to ligate the hairpin cassette

to a binary vector.

Vectors with both sense and antisense fragments of target genes ligated were sequenced as

described in Section 3.3.5. The primers used for sequencing were SIntron and ASIntron in

separate reactions. Sequencing results were analysed using FinchTV 1.4.0 (Perkin Elmer, Inc.)

85

and MultAlin alignment software (Corpet 1988). After confirming the correct orientation of the

gene fragments, the hairpin expression cassette was transformed into the binary vector pART27.

4.3.2 Cloning into the binary vector pART27

The binary vector pART27 was used for plant transformations (Gleave 1992). The hairpin

cassette from pCleaver for the nematode genes, gfp and gene-free control were digested out

using NotI and ligated to NotI linearised, dephosphorylated pART27. Antarctic phosphatase

(New England BioLabs) was used for dephosphorylation. Five units of enzyme per 1.5 µg of

NotI digested vector DNA was incubated at 37 °C for 30 minutes followed by deactivation at 65

°C for 5 minutes. Ligated vectors were transformed into E. coli JM109 as described in section

3.3.4 and cultured on selective media containing LB and 100 µg/mL of spectinomycin

dihydrochloride. Colony PCRs were done using the primers 35SART (5′-GTCTTGATGAGAC

CTGCTGCGTA-3′) and SP6 (5′-CATACGATTTAGGTGACACTATAGA-3′). The 35SART

primer binds in the 35S promoter sequence of the hairpin cassette while SP6 binds near the right

border of the vector pART27 (Figure 4.2).

A confirmatory digest of cloned, purified plasmid DNA was done with the enzyme SalI to

confirm the presence of insert. Confirmed plasmid DNA was then used to transform

Agrobacterium tumefaciens for plant transformations.

Figure 4.2: Map of the vector pART27. Primer binding sites and restriction enzyme sites are

indicated.

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4.3.3 Agrobacterium tumefaciens transformation

Agrobacterium tumefaciens strain GV3101 was prepared using the calcium chloride method

(Sambrook and Russell 2001). Briefly, 250 µL of overnight culture of A. tumefaciens was used

to inoculate 100 mL LB broth in a 500 mL flask. The culture was incubated on a shaker at 28°C

in the dark until OD600 reached 0.8 after which the flask was chilled on ice for 10 minutes. The

culture was transferred to 50 mL centrifuge tubes and centrifuged at 4 °C for 30 minutes at

3,000 g. Supernatant was discarded and pellet was resuspended in 1 mL of sterile 20 mM

calcium chloride at 4 °C. Aliquots of 50 µL were dispensed in pre-chilled eppendorf tubes, snap

frozen and stored at -80 °C. All transformations were done with 50 µL of competent cells and

500 ng of plasmid DNA using the heat shock method. Briefly, competent cells were thawed on

ice and vector DNA was added followed by a further incubation of 20 minutes on ice. The cells

were then incubated at 37 °C for 5 minutes in a water bath. One mL of LB was added and

incubated for 3 hours in the dark at 28 °C with shaking (200 rpm). After 3 hours, an aliquot

(150-200 µL) of this culture was plated on LB media containing 100 µg/mL of spectinomycin

dihydrochloride and 25 µg/mL rifampicin. Bacteria were allowed to grow for 2 days at 28 °C

after which colonies were picked and resuspended in 20 µL PCR water. After initial failure to

obtain an amplicon using 5 µL of this bacterial suspension as template in PCR, the same

suspension was heated at 96 °C for 10 minutes prior to adding the PCR reaction master mix.

35SART and SP6 primers were used for these PCRs with the following cycling conditions:

initial denaturation at 96 °C for 2 mins; 25 cycles of 96 °C for 30 secs, 55 °C for 30 secs and 72

°C for 30 secs and an extension step at 72 °C for 7 mins. Amplicons were visualised on a 1%

agarose gel. Positive colonies were cultured in 5 mL LB broth supplemented with 100 µg/mL of

spectinomycin dihydrochloride and 25 µg/mL rifampicin.

4.3.4 Plant transformation

Arabidopsis thaliana ecotype Columbia-0 was used for transformation using the floral dip

method (Bent et al. 2006). Arabidopsis seeds were grown in the glasshouse under controlled

conditions (22 ± 5 °C and 16 hours light) in trays. The soil mix was prepared with 1 part

Murdoch mix (composted pine bark: coarse river sand: coco peat in a 2:2:1 ratio) and 1 part

Richgro seed & cutting mix with fertilizers and minerals i.e. Grower’s blue (60 g), Osmocote

(60 g), dolomite (20 g) and calcium carbonate (15 g) added per 40 L of soil mix. After

germination, the plants were transferred to plastic cups (200 mL) with holes at the base. First

flowering heads were cut off to encourage growth of multiple heads which were then subjected

to floral dip transformation.

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To prepare A. tumefaciens inoculum, 100 mL of LB supplemented with 100 mg/L

spectinomycin dihydrochloride and 25 mg/L rifampicin was inoculated with 2 mL of a stock

bacterial culture and incubated at 28 °C in the dark until the OD600 was 0.1-0.2. Bacterial cells

were pelleted by centrifugation and resuspended in an equal volume of 5% sucrose. Ten to

fifteen plants for each construct were used for transformation. Silwet-77 surfactant was added at

a concentration of 0.05% before dipping florets into this solution for 10-15 seconds. Plants were

covered with clear plastic for one day to maintain humidity and then grown and watered as

normal until siliques were matured and dry. At this stage, watering of the plants was stopped

and brown paper bags were used to cover the siliques. Completely dried siliques were cut off at

the base of flowering stems and stored in paper bags until threshed. After threshing, seeds were

passed through a 2 mm sieve and collected in 5 mL plastic tubes for storage.

4.3.5 Screening for Transgenic (T1) plants

Seeds collected from transformed plants were sterilised by the vapour-phase surface sterilisation

method (Bent et al. 2006). Seeds were transferred to 2 mL microfuge tubes to about 50 µL

packed volume and placed in a rack upright with lids open in a desiccator in the fumehood. At

the centre of the desiccator was a glass bottle containing 100 mL of 12.5% sodium hypochlorite.

Four mL of hydrochloric acid was slowly poured into the glass container after which the

desiccator lid was closed. After 12-14 hours, the lid was slightly opened in the fumehood to

allow gas to escape for 20 minutes. The rack was then transferred to a laminar flow cabinet and

the tubes left open for 30 minutes to remove residual fumes.

Seeds were then spread with 0.4% agar on MS media plates (1/2 Murashige and Skoog basal

medium with Gamborg vitamins, 3% sucrose, 0.8% Agar) supplemented with 50 µg/mL of

kanamycin monosulfate to screen for transformed plants. After initial incubation at 4 °C in the

dark for two days to break dormancy, the plates were placed in a growth chamber (Conviron

ATC40) with controlled conditions (16 hour light @150 µmol/m2/s, 40% RH and 23 ± 2 °C).

After screening for at least three weeks, plants that survived were transferred to 200 mL plastic

cups containing soil mix. These plants were grown in the same growth chamber. Once the plants

just started to bolt, ‘Aracons’ with transparent plastic tubes (height: 60 cm) with holes for

ventilation were placed in every cup to prevent cross fertilisation and guide dried seeds to fall

into the Aracon container. When bolts and pods were completely dry, seeds were collected by

cutting the bolts at the base and inverting the Aracon assembly into a paper bag. Seeds were

sieved and collected into 5 mL sterile plastic tubes.

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4.3.6 Screening and challenge of T2 plants

Seeds from T1 plants packed in 50 µL volume were sterilised as described previously. They

were then spread on MS medium plates as described for T1 plants and incubated at 4 °C for two

days. A control plate spread with wild-type seeds on medium with kanamycin was also

prepared. Seedlings were allowed to grow at optimum growth conditions (23 °C and 16 hour

light) until plantlets on control plates were dead, and clear selection was observed for transgenic

plants.

Plants were then transferred to 56-well trays (well capacity: 65 cm3) containing soil and placed

in the growth chamber (23 ± 2 °C, 16 hour light @150 µE/m2/s and 40% RH). After initial

acclimatisation of plants in soil and growing for another week, each plant was infected with 200

active J2s of M. incognita extracted from tomato plant roots.

4.3.7 Nematode collection for infection

J2s of M. incognita were extracted from roots of heavily infested tomato plants (cv. Grosse

Lisse) and the plant’s rhizosphere, using a mist apparatus as described by Tan et al. (2013).

Briefly, two layers of coffee filters containing root and soil material were placed on a sieve in

funnel and sprayed for 10 seconds with a fine mist every 10 minutes for two days. Nematodes

which passed through the filters to the lower part of the mist apparatus in the tubing were

collected in a 240 mL glass bottle.

The bottle was placed on lab bench for 4-5 hours to allow the nematodes to settle. Excess water

was removed using a vacuum pump and the nematodes cleaned using the sucrose gradient

method (Amin et al. 2014). Sucrose gradient columns were made with 3 mL of 50% sucrose

solution at the bottom carefully overlaid with 4 mL of 10% sucrose solution in a 14 mL tube.

Nematodes suspended in water (4 mL) were carefully poured on top of the column and

centrifuged for 15 minutes at 2,150 g with slow deceleration so as not to disturb phases.

A layer of clean J2s was formed at the junction of 50% and 10% sucrose solutions, which was

carefully pipetted out and poured onto a 10 µm filter. One litre of sterile water was passed

through this sieve to wash sucrose off the J2s, which were then collected for plant infection.

4.3.8 Nematode infection of transgenic plants

Two hundred active J2s were used to infect transgenic Arabidopsis plants (4 to 10 events for

each target gene) by pipetting water containing nematodes on four sides of each plant root ~1

inch deep in the soil. Fourteen uniform replicate plants were infected for each transgenic event

and 20 for wild-type plants. Plants were watered sparingly for two days following nematode

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infection. Four weeks after infection, plants were uprooted and roots were thoroughly washed to

eliminate soil particles. Galls were counted for 10-11 plants per transgenic event (16 for wild-

type) using a dissecting microscope. Roots separated at the collar were dried in an oven for 16-

18 hours at 55 °C and weighed afterwards. The number of galls per gram dry root weight was

calculated for each plant.

4.3.9 Nematode development after infection

Three to four plants for each transgenic event were uprooted after five weeks of infection and

females were dissected out of the galls. These were stained with acid fuschin to study

morphological differences between nematodes feeding on transgenic plants and those on

controls. After collecting females in water in a 1.5 mL microfuge tube, they were centrifuged

for 30 sec at 500 g. Water was removed by pipetting and 200 µL acid fuschin staining solution

was added. The tube was incubated at 95 °C for 2 minutes with gentle agitation and centrifuged

again at 500 g for 30 seconds to remove the staining solution. Stained females were washed 2

times with water and viewed on a glass slide under bright field view (Olympus BX-51

microscope) with 4x and 10x magnification.

4.3.10 Statistical analyses

Analysis of variance (ANOVA) was done for comparison between treatments and calculation of

means, standard deviation and standard error in the statistical software package SPSSv20 (IBM

Corporation, US) Significance between treatments and pair-wise comparisons were done using

Tukey’s test at a significance level of 0.05. Microsoft Excel Analysis ToolPak was used for

construction of bar charts.

4.3.11 Confirmation of T-DNA insertion

Four transgenic events per gene were selected for PCR to confirm integration of target DNA

into the Arabidopsis genome. Leaf samples were collected from three plants of each event for

DNA extraction using the CTAB method. About 100 mg of plant sample was transferred to 1.5

mL microfuge tube with three 3.2 mm stainless steel beads (Qiagen, Australia). Tubes were

frozen in liquid nitrogen and the leaves ground to fine powder using a Tissue Lyzer (Qiagen,

Australia) at 20 Hertz for 2 minutes. One mL of pre-heated (65 °C) CTAB buffer (2% CTAB,

100 mM Tris-HCl, 20 mM EDTA, 1.4 M NaCl, pH 5.0 and freshly added 1% PVP and 2% 2-

marcaptoethanol) was added to each tube and vortexed for 2 minutes. Tubes were incubated at

65 °C for 30 minutes and then centrifuged at 14,000 g for 2 minutes. Supernatant was collected

in a 2 mL microfuge tube and equal volume of chloroform:isoamyl alcohol (24:1) was added,

the emulsion vortexed for 1 minute and then centrifuged for 2 minutes at 12,000 g to separate

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phases. The aqueous layer was collected in a clean microfuge tube, 1/10th volume of 7.5 mM

Ammonium acetate and 2 volumes of 100% ethanol were added and precipitated overnight at 4

°C, after which the tubes were centrifuged at 16,000 g for 15 minutes to pellet the nucleic acids.

The supernatant was discarded and the pellet washed with 70% ethanol, dried and later

resuspended in 100 µL of sterile water.

Resuspended DNA was quantified using a Nanodrop spectrophotometer, and 100-200 ng of this

DNA was used in standard 20 µL PCR reactions. Primers used were NptII-F (5′-TGCTCCTGC

CGAGAAAGTAT-3′) and NptII-R (5′-AATATCACGGGTAGCCAACG-3′) to amplify a 364

bp fragment of the nptII gene in the binary vector. Amplicons were visualised on a 1% agarose

gel.

4.3.12 Confirmation of transcription of nematode silencing signals

RNA extraction was done with the Trizol method and treated with DNaseI as described

previously for nematodes in Section 3.3.2, except that 1 mL of diluted Trizol LS reagent was

used. After drying, the pellet was resuspended in 80 µL of sterile water from which 500 ng was

used to synthesise cDNA using the High capacity cDNA synthesis kit (Applied Biosystems) in a

20 µL reaction. One in ten dilutions of this cDNA were used in 20 µL PCR reactions using the

nematode gene specific primers with the following cycling conditions: initial denaturation at 96

°C for 5 mins; 30 cycles of 96 °C for 30 sec, 55 °C for 30 secs and 72 °C for 1 min and an

extension step at 72 °C for 7 mins. Amplicons were visualised on a 1% agarose gel.

4.4 Results

4.4.1 Analysis of transgenic plants

After floral dip of Arabidopsis plants, the seeds were screened for selection of transformants.

The transformation efficiency varied for the different constructs used. Table 4.1 shows the

number of events that survived kanamycin screening for three weeks for both T1 and T2

generations. In the second screening some lines appeared to be non-transformed, and none of

the selected transformants survived for the genes ego-1, alg-1, 2242, rol-6 or the null vector.

The genes in the RNAi pathway are conserved genes across eukaryotes and there were

mismatches between the M. incognita gene sequences used for transformation and those of A.

thaliana, but there were no continuous matches of 18-21 bp for any of the genes studied. No

phenotypic differences were observed for the plants transformed with the nematode sequences

used or with gfp. Healthy dry seeds were produced by all events (T1 and T2). Screening of T1

plants on plates and plants growing in cups with the Aracon assembly are shown in Figure 4.3.

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Table 4.1: The number of transgenic events of A. thaliana that survived after kanamycin

screening. All of the harvested seeds were screened at the T1 generation, while for T2 a

maximum of 10 events per construct were screened.

M. incognita

gene

No. of T1

transformed

events

No. of T2

transformed

events

M.

incognita

gene

No. of T1

transformed

events

No. of T2

transformed

events

dcr-1 23 9 rha-1 18 10

drh-3 14 10 alg-1 3 0

vig-1 8 4 2242 0 -

ego-1 3 0 rol-6 1 0

mut-7 8 5 gfp control 13 9

drsh-1 12 10 Null vector 4 0

pash-1 6 5

(A) (B)

Figure 4.3: Kanamycin screening of T1 transgenic events. (A) Transformed events (T1)

growing on kanamycin selection for three weeks. (B) Plastic cups with one transgenic event

(T1) in each, with an Aracon assembly to prevent cross fertilisation and easy seed collection.

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Screening for T2 generation plants revealed that there must have been some false positives

among selected T1 events, since the following putative events did not yield viable plants when

selected on kanamycin a second time: ego-1, alg-1, 2242, rol-6 and null vector.

After the second screening, the T2 events that grew well and were used for nematode challenge

were transformed with the constructs targeting dcr-1, drh-3, vig-1, mut-7, drsh-1, pash-1, rha-1

and gfp vector. Since this generation was still segregating, a 3:1 ratio of transformed to non-

transformed lines was evident on some plates (Figure 4.4). Wild-type seeds on control plates did

not survive the kanamycin screening. Antibiotic screening and plants growing in trays after

infection are presented in Figure 4.4.

4.4.2 Analysis of nematode infection

Between 10-11 plants were analysed for infection from each transgenic line. With as many as an

average of 2,300 galls per gram of dry root weight, wild-type plants were clearly highly

susceptibility to infection by M. incognita. There was no significant difference in the average

number of galls/g dry root between the gfp transgenic events analysed (p<0.05). Also infection

levels were not significantly different from the infection levels on wild-type control plants

(Figure 4.5).

From the dicer complex group of genes, transgenic events were analysed for dcr-1 and drh-3.

Significantly reduced infection was observed for all nine hpdcr-1 events when compared to

wild-type or hpgfp events (p<0.05). For five out of the nine events there was reduced nematode

infestation, by 50% or more, with the greatest reduction of 71% (event 8) (Figure 4.5). The

lowest reduction in infestation was 35% for hpdcr-1 events. Similarly, there was significantly

reduced infestation in all events of hpdrh-3 plants (p<0.05). However, the range of reduction in

(A)

(B)

(C)

Figure 4.4: Kanamycin screening of T2 transgenic plants. (A) Antibiotic screening of T2 plants. (B)

Control plate with wild-type seedlings that did not survive kanamycin screening. (C) Transgenic

plants growing in 56-well trays after nematode infection.

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infection was better than dcr-1 events: ranging from 42-74% and with seven of the ten events

showing reduced nematode infection by 50% or more while three were 70% less than controls.

All transgenic events expressing hpRNA corresponding to the gene drsh-1 and its co-factor

pash-1 reduced nematode infection significantly (p<0.05). The infection reduction for the 10

events of drsh-1 was between 30-89%: eight of the events exhibited a reduction of >50%. This

group of transgenic events also showed most variation for the average number of galls/g dry

root between events. The five events expressing hppash-1 were all significantly less infested

than controls, by as much as 64%. The highest infested line from this group had 30% fewer

galls/g dry root than controls (Figure 4.5).

Two other groups of RNAi effectors were targeted through host-delivered silencing signals. The

RISC effector vig-1 when targeted reduced infection significantly (p<0.05). Four lines tested for

nematode infection showed a reduction in infection of 38-64% compared to controls (Figure

4.5). Five transgenic events for the nuclear RNAi gene mut-7 were also tested for nematode

susceptibility. Four of the events showed significantly reduced infection compared to controls,

while one of the events (event 3) was not significantly different because of the variation in

replicates even though the infection reduced by 38%. The transgenic events targeting nuclear

RNAi effector rha-1 also showed significantly reduced infestation for 9 of the 10 events tested.

The reduction in plant infestation targeting this gene was in the range of 33-69% (Figure 4.5).

With the exception of two (one for mut-7 and one for rha-1), transgenic events carrying hp of all

7 genes studied had significantly reduced nematode infestation compared to the controls

(p<0.05). Sequences from genes of the microprocessor complex and dicer complex appeared to

be the most effective in reducing nematode infestation through HIGS. Overall, strongest

reduction in infestation of the T2 generation of A. thaliana plants was an 89% reduction in

nematode infestation. This correlates quite well with results from soaking experiments.

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(A) (B)

(C) (D)

(E) (F)

(G) (H)

Figure 4.5: Nematode infection, as measured by the number of galls per gram dry weight of root

tissue, compared to controls (A) hpgfp events, (B), hpdcr-1 events, (C) hpdrh-3 events, (D) hpdrsh-1

events, (E) hppash-1 events, (F) hpvig-1 events, (G) hpmut-7 events and (H) hprha-1 events.

Comparisons are based on average number of galls/ gram of dry root weight for replicate plants (n =

9 to 16) between wild-type and transgenic events (means ± standard error). Bars having different

letters indicate significant differences (p<0.05).

95

4.4.3 Female morphology

Adult female nematodes were dissected from plant roots five weeks after infection. Three events

for each hpRNAi vector were selected for this. The females extracted from transgenic plants

exhibited three abnormal phenotypes when compared to controls. These were smaller size,

elongated (fusiform) body shape and a transparent, sickly appearance. The females extracted

from plants with hpRNA targeting the dicer complex and miRNA processing genes were

smaller in size than those extracted from control wild-type and hpgfp transgenic plants. This

small size of adult female nematodes was most prominent for drh-3, drsh-1, pash-1 and rha-1

targeting plants, where nematodes were smaller in diameter at the posterior end compared to

saccate large females produced in roots of wild-type and hpgfp plants (Figure 4.6).

The types of changes in adult nematode morphology are shown in Figure 4.6 for nematodes

feeding on transgenic plants targeting drh-3, pash-1, mut-7 and rha-1. The highest percentage of

nematodes with modified phenotypes was found for pash-1 targeting lines in which 65% of the

extracted females were fusiform or elongated. The percentage was 62% for rha-1 targeted

nematodes, and for targets against drh-3 and drsh-1 the elongated phenotype was 56% and 46%

respectively.

The transparent appearance of nematodes was observed for those feeding on plants producing

dcr-1, drsh-1, pash-1 and vig-1 silencing signals. Clearly these nematodes did not make egg

masses and had impaired reproduction. Fusiform and transparent phenotypes were also evident

for nematodes infecting plants producing targets against vig-1 and pash-1. Although the

percentage of nematodes with transparent bodies was low, only about 5% for vig-1 and pash-1

treatments, nevertheless none of the nematodes growing on control plants showed this

phenotype.

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hpgfp hpgfp Wt Wt

hpdcr-1 hpdrh-3

hpdrsh-1 hppash-1 hpvig-1

hpdcr-1 hpdcr-1 hpdrh-3

hpdrsh-1 hppash-1

hpvig-1 hpmut-7 hpmut-7 hprha-1 hprha-1

4.4.4 T-DNA insertion and dsRNA transcription

A 364 bp fragment of the nptII gene was amplified from DNA extracted from different

transgenic lines. Four events displaying significantly lower infection (p<0.05) for each

transgene were chosen for PCRs and RT-PCRs. Figure 4.7A shows the bands amplified from

the DNA of all the events tested, which confirmed the transgenic status of the plants and

successful T-DNA insertion in the plants challenged with nematodes.

To confirm RNA transcription in the same events, RT-PCRs were done with gene specific

primers on the plant-derived RNA. Amplified bands are shown in Figure 4.7B. The intensity of

the bands was different for different transgenic events. The specific nematode genes could not

be amplified from two of the transgenic lines i.e. for one line each of dcr-1 and pash-1.

Figure 4.6: Adult females dissected from wild-type (Wt) and hpRNA transgenic A. thaliana

plants five weeks after infection. Nematodes were stained with acid fuschin. Scale bar:

500µm.

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4.5 Discussion

In this study, A. thaliana plants were successfully engineered to constitutively express dsRNA

targeting 7 different genes affecting the RKN M. incognita. HIGS in this study resulted in a

significant reduction in infection for all targets, and also defects in nematode development for

some targets. There were no apparent phenotypic effects in any of the transgenic Arabidopsis

plants, although there could be other less obvious changes. There were no significant

differences in the level of infection between wild-type and dsgfp producing control plants,

indicating that there was no effect on nematode parasitism of plant transformation in itself. In

contrast, all of the host-delivered triggers significantly impaired the ability of nematodes to

infect, produce galls and reproduce on transgenic plant roots.

One aspect considered at the start of RNAi studies for nematode control was a concern that the

size of dsRNA molecules might prevent their uptake. However, this does not seem to be a

problem, since M. incognita J2s have been shown to be able to take up 800 bp dsRNA targeting

the cathepsin L-cysteine gene from solution, and from plants they are able to take up 28 kDa

GFP protein (Urwin et al. 1997b; Shingles et al. 2007). In this study, the size of the dsRNA

sequences used to generate transgenic lines was between 187-669 bp, and the results clearly

indicate that dsRNA had been taken up, since all treatments reduced nematode infestation

significantly. Different band intensities of amplified DNA (PCR and RT-PCR) possibly point to

different levels of hpRNA produced in different transgenic lines. This aspect can be investigated

further by analysis of gene copy number and quantitative transcript analysis of homozygous

lines of the next generation of these plants.

Resistance to nematode infestation varied with the gene being targeted through HIGS. There

were also differences in responses to different events of the same transgene. This is a common

phenomenon encountered with different events for most transgenic plant experiments. There are

various factors which cause this variation, and these include gene copy number of inserts, the

site of insertion, and flanking sequences, and all influence the level of transgene expression,

M hpdcr-1 hpdrh-3 hpvig-1 hpmut-7 hpdrsh-1 hppash-1 hprha-1 hpgfp

A

B

Figure 4.7: (A) Amplified nptII gene fragment from genomic DNA of four events for each hpRNAi

vector. (B) Amplified nematode-specific transcript from transgenic lines using RT-PCR. M = 100bp

DNA ladder.

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which translates into different efficiencies of events to amplify the silencing triggers, and

deliver them via the feeding tube to the feeding nematode. An additional factor which may

increase the standard deviations observed may be the presence of both homozygous and

heterozygous plants in the analysed material. Unless, a gene copy number analysis is done for

identifying homozygous single copy insert plants, it is difficult to rank the target genes

confidently in order of effectiveness. Nonetheless, the events tested still reduced nematode

infection success significantly, and the range of responses for different events of the same gene

reflects some of the factors discussed above.

The nematode genes targeted in this study are involved in the endogenous gene regulation

system which may affect nematode developmental pathways. A similar transparent appearance

following host-delivered RNAi has been reported before by Yadav et al. (2006) who suggested

it was caused by a lack of gut granules. However this is interpreted, it is clear that such

nematodes are extremely unhealthy. It is also significant that developing eggs can normally be

seen inside the female body, but this was not the case for the transparent females. Therefore,

reproduction was either non-existent or substantially reduced for these females. Similarly,

nematodes exhibiting a fusiform (elongated) body may result from delayed growth or defective

moulting. Challenge of several generations of nematode on these plants will give a good idea on

the magnitude of such effects on nematode development and reproduction, and also provide

information on whether such effects are stable or amplified through generations.

In considering what genes might be appropriate to use as targets for nematode control via HIGS,

evidence is now accumulating, at least for RKNs, that many different types of targets can be

chosen. Other work has tended to focus on genes involved in parasitism in relation to nematode

entry and migration in root tissues, avoiding host defences, or genes required for giant cell

induction and function. In the work undertaken here, it is clear that apart from genes involved in

parasitism and development, those involved in gene regulation such as the RNAi pathway are

also suitable target genes for nematode control, especially since down-regulation is likely

directly or indirectly to affect a wide range of biochemical and metabolic pathways. Similarly,

highly deleterious effects have been reported for genes involved in mRNA metabolism for both

in vitro soaking (95% lethality) and host-delivered silencing (88% reduced infection) in RNAi

studies on H. glycines (Alkharouf et al. 2007; Klink et al. 2009).

It is interesting to point out that drsh-1 targeting plants impaired nematode infectivity by more

than its co-factor pash-1, but when nematode development was analysed, RNAi of pash-1

seemed to have more deleterious effects than that of drsh-1. This suggests that a combinatorial

host-delivered knockdown approach for these genes might result in higher impact, disrupting

both infectivity and development of RKNs.

99

The seven M. incognita genes transformed into plants in this study show high similarity to M.

hapla orthologous sequences. This homology was between 90-94% with stretches (>25 bp) of

sequences matching perfectly except for vig-1. With this degree of homology in common target

genes, it is possible that the transgenic plants generated for resistance to M. incognita could also

be resistant to M. hapla. Such cross-resistance is entirely possible since plant-derived RNAi of

Mi16D10 reduced egg production per gram root for four species of RKN (M. incognita, M.

javanica, M. arenaria, and M. hapla), by between 69% and 93% (Huang et al. 2006).

Another possibility also exists that using data from transgenic plants to identify the most

effective targets for silencing in root-knot nematodes: it may be possible to deliver dsRNA

stabilised in some way and sprayed onto plants. This approach would avoid the issues of

acceptance of transgenic plants, and could well be applied to control of other above ground

pests through topical dsRNA application. Some success from this approach has been reported

for pests like the diamondback moth and citrus psyllid (Gong et al. 2013; Killiny et al. 2014).

Knowledge gained from experiments using Arabidopsis on control of PPNs should be

translatable for application in commercial crops for example for soybean, HIGS reduced the

damage caused by soybean cyst nematode by up to 67% (Matthews et al. 2014). An important

aspect in relation to commercial application is the selection of transgene sequence used, which

should have absolutely no off-targets, either for the host plant or non-target organisms. It is only

really possible to ensure that this is the case once full genomic sequences are available for the

crop plant of interest, and for other species that might encounter that crop. Nevertheless, for

obligate parasites such as PPNs, RNAi technology is a powerful tool for reverse genetics

studies, and can be used to probe mechanisms underlying parasitism and development.

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Chapter 5

The effects of RNAi treaments with different regions of

the Dicer-like gene on the viability, parasitism and

reproduction of M. incognita

101

The effects of RNAi treaments with different regions of the Dicer-like gene

on the viability, parasitism and reproduction of M. incognita




5.1 Abstract

RNAi is a gene regulation mechanism which has homologous functional genes in eukaryotes.

The ribonuclease Dicer-like enzyme plays an essential role in RNA recognition and processing

of dsRNAs into siRNAs/miRNAs, which act downstream in the sRNA pathways. The only

dicer-like gene of nematodes, dcr-1 has been characterised in C. elegans but not in parasitic

nematodes. In this study the aim was to down-regulate the expression of the dcr-1 of M.

incognita, targeting seven different mRNA regions of this gene separately, to investigate the

effects on nematode infectivity and reproduction. The pattern of expression of dcr-1 in the

nematode and the effect of its down-regulation on other genes of the pathway were also

assessed. The dcr-1 expression was detected predominantly in the intestine and body wall,

whilst the level of reduction in host plant infection and reproduction depended on the region of

target mRNA chosen for silencing, with most reduction found for the region closest to the 5′ end

of the gene which also showed down-regulation of gene expression. Different triggers also had

differential effects on subsequent gene expression, as assessed by quantitative RT-PCR. The

effects were not linked to GC content of trigger or target secondary structure, although low

positional entropy and high base-pair probability was detected for target gene sequences which

were less effective. The in vitro RNAi of dcr-1 in these experiments led to altered expression of

other genes of the pathway and reduction in nematode infection and reproduction.

Keywords: Dicer, RNAi pathway, plant parasitic nematode, target mRNA region, in vitro

RNAi.

5.2 Introduction

Processing of dsRNA into siRNAs is carried out by the endonuclease enzyme dicer. There are

different classes of dicer-like genes in different organisms, but it seems to be an essential feature

of the RNAi pathway. It is regarded as the convergence point for various sRNA pathways in

nematodes. Dicer-1 has been characterised in detail in C. elegans using mutants, and is known

to be involved in generating ~22-23 bp siRNAs, and in developmental timing and innate

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immunity (Ketting et al. 2001; Iatsenko et al. 2013). It has also been studied in detail in Giardia

intestinalis, including its protein domain architecture, crystal structure and mechanism of action

(MacRae et al. 2006). Post-translational modification of the dicer gene product has been

demonstrated in C. elegans and humans, and this has been suggested as a requirement for

division of labour in the small RNA pathways (Sawh and Duchaine 2013). However, the dicer

gene/complex has not been studied in anywhere near the same detail in any PPN.

The efficiency of dsRNA mediated gene silencing appears to be affected by the region of target

gene transcript chosen for synthesis of dsRNA (Sukno et al. 2007; Arguel et al. 2012). The dcr-

1 gene in nematodes is relatively large, and is made up of a series of different protein domain

coding regions. In the experiments described here, the effects of using sequences from different

domains of the dcr-1 of M. incognita as triggers in silencing this gene was examined. The

expression pattern of M. incognita dcr-1 and effects of its silencing on RNAi effectors was also

studied. Subsequent ability of treated nematodes to parasitise plants and to reproduce was

reduced indicating the importance of dicer gene for development of the nematode.


5.3.1 Sequence analysis and primer design

C. elegans Dicer-1 sequence was retrieved from wormbase (www.wormbase.org) and used to

identify a genomic contig (CABB01000157) of M. incognita from NCBI as described in

Chapter 2. Seven primer pairs were designed to amplify sequences potentially encoding the

seven protein domains identified previously (Chapter 2). Primer sequences, their position on the

genomic contig and their respective amplicon lengths are indicated in Table 5.1.

Table 5.1: Primers used to amplify seven protein domain coding regions of M. incognita dcr-1

gene. Domain names and amplicon sizes are indicated.

ID Primer 5′− 3′ Flanked

Domain

Amplicon

(bp)

Position on

contig

MiDcr1HelATPB-F

MiDcr1HelATPB-R

TTGATGTAACAACCTCTGGGA

AATTCAAGGTTGAATTACTTGATCG

DEAD-like

Helicase

125 19818-19945

MiDcr1HelCT-F

MiDcr1HelCT-R

CTTGAATAAATGAACGAAAGTCCATC

AGACAGGCGATGAAGATAGG

Helicase C

terminal

181 17995-18177

MiDcr1DSRBF-F

MiDcr1DSRBF-R

TTCCTGCAGGCAAAAGATTGTC

GGTACTGTGCAAAATTACCATCTG

Dicer Dimer 251 17326-17519

MiDcr1PAZ-F

MiDcr1PAZ-R

CATTCTAGCAGATGTATGGTCAAC

GAATCTCCAACTACACCTTCAGATG

PAZ 272 16037-16182

MiDcr1RiboC1-F

MiDcr1RiboC1-R

CATGTGGATCGAATTTAAGTGTTTC

CATTGACAACTTCGAGTGCTG

Ribonuclease

III

228 13597-13881

MiDcr1RiboC2-F

MiDcr1RiboC2-R

ACAGAAATTAACAAACTTTTCGACC

TTAGCGTTTGGAATTCCTTGG

Ribonuclease

III

233 12434-12738

MiDcr1DSRM-F

MiDcr1DSRM-R

TCCAATTTTCTTAAATGTTGAAGTGC

TTGGAACGAAAATTAGAAACAGGC

Double

Stranded RNA

binding domain

139 11897-12039

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5.3.2 Cloning and synthesis of dsRNAs

Restriction enzyme sites for XhoI and KpnI were added to the primers and M. incognita cDNA

from mixed stages was used as template to amplify gene fragments as described previously

(Section 3.3.3). Amplicons were purified from 1% agarose gel using the Wizard® SV Gel and

PCR Clean-Up system (Promega, Australia) according to manufacturer’s protocol and

quantified with a Nanodrop spectrophotometer. The seven gene fragments were then cloned into

the transcription vector pDoubler after ligation. Transformation with E. coli and PCR colony

screening was done as described in Section 3.3.4. Plasmid purification and sequencing were

done as described in Section 3.3.5 using the primers M13-F and M13-R in separate reactions.

DNA template for dsRNA synthesis for all gene fragments and gfp for use as control was

generated using PCR with the primer T7 (5′-TAATACGACTCACTATAGGG-3′) in a standard

20 µL reaction. After clean-up, 1 µg of each template was used to synthesise dsRNA

corresponding to gfp (dsgfp) and the seven dcr-1 regions: DEAD-like helicase (dsD1), helicase

C terminal (dsD2), dicer dimer (dsD3), PAZ (dsD4), the two ribonuclease III domains (dsD5

and dsD6) and double stranded RNA binding domain (dsD7) using HiScribe T7 in vitro

transcription kit (New England BioLabs). Synthesised dsRNA was digested with DNase I and

run on 1% agarose gel after purification to assess integrity.

5.3.3 In situ hybridisation

In situ hybridisation of mRNA was done to study expression pattern for M. incognita dcr-1

according to the protocol described by De Boer et al. (1998) using the DIG labelling and

detection kit (Roche Life Science). Probe templates (sense and antisense Ribonuclease III

coding region RiboC1) were digested from pDoubler using the enzymes XhoI/NotI and

KpnI/NotI and one microgram of this template was used in separate T7 initiated transcription

reactions for probe synthesis. In addition to the ribonucleotides ATP (10 mM), CTP (10 mM),

GTP (10 mM) and UTP (6.5 mM), digoxigenin-11-UTP (3.5 mM) was added in the reaction to

label single stranded RNA and incubated at 37 °C for 8 hours. RNA was purified by lithium

chloride precipitation and ran on 1% agarose gel to assess integrity.

Freshly hatched J2s of M. incognita packed in a 30 µL volume were collected in a 1.5 mL tube

by centrifugation at 1,200 g for 2 minutes and fixed in 2% paraformaldehyde in M9 buffer for

18 hours at 4 °C, followed by an incubation of 4 hours at 22 °C. Nematodes were pelleted,

pipetted on to a glass slide and cut into sections using a 32 gauge syringe needle. After

collecting nematode sections in the tube by washing with M9 buffer, they were digested with

proteinase K (0.5 mg/mL in M9 buffer) at 22 °C for 20 minutes. After removing the solution,

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nematodes were frozen on dry ice and incubated for 1 minute each in methanol and acetone

respectively followed by rehydration at 22 °C for 20 minutes in 20% acetone.

At this stage, nematodes were divided equally into two tubes and pre-hybridised for 1 hour at 50

°C in the hybridisation buffer which contained 50% deionised formamide, 4x SSC, 2% SDS,

1% blocking reagent, 0.1x maleic acid buffer, 0.2 mg/mL Ficoll 400, 0.2 mg/mL PVP, 0.2

mg/mL bovine serum albumin, 1 mM EDTA, 0.2 mg/mL fish sperm DNA and 0.15 mg/mL

yeast tRNA. Nematode sections were hybridised with the riboprobe (300 ng/mL) in fresh

hybridisation buffer at 50 °C overnight in separate reactions for sense and antisense probes.

J2 sections were then washed three times with 4x SSC at 50 °C for 15 minutes each followed by

a 10 minute wash at 37 °C with NTE buffer (0.5 M NaCl, 10 mM Tris-HCl, 1 mM EDTA; pH

8.0). Un-hybridised probe was digested using RNase A (60 µg/mL in NTE buffer) at 37 °C for 1

hour followed by three washes at 50 °C with 0.1x SSC, 0.1% SDS. Detection of hybridisation

was done at 22 °C, where nematodes were washed in maleic acid buffer and incubated in 1%

blocking solution in maleic acid buffer. After labelling for 2 hours in alkaline phosphatase anti-

digoxigenin Fab fragments diluted (1:1000) in blocking solution, nematodes were washed three

times (15 minutes each) with maleic acid buffer with Tween-20 (0.05%). Following another

wash with detection buffer, nematode sections were stained with nitroblue tetrazolium (337

µg/mL) and 5-bromo-4-chloro-3-indolyl phosphate (175 µg/mL) in 250 µL of detection buffer

without agitation at 5 °C overnight. Staining was stopped by washing with water two times and

nematodes were observed with a microscope (Olympus BX-51) under bright field view.

5.3.4 DsRNA soaking

Seven thousand freshly hatched M. incognita J2s were fed with dsRNA derived from each of

the seven dicer fragments. Dsgfp and no dsRNA reactions were set up as controls. FITC (1

mg/mL) was added in a separate reaction to monitor solution uptake. Reactions were composed

of M9 buffer with 1 mg/mL dsRNA, 50 mM octopamine, 3 mM spermidine, and 0.05%

gelatine. Nematodes in soaking solution were incubated at 25 °C for 16 hours.

5.3.5 RNAi phenotypes, RNA extraction and plant infection

After 16 hours of incubation in dsRNA, nematodes were harvested from the solution, washed

with clean water and observed under a microscope as described previously (Section 3.3.8). The

soaking solution was run on a 1% agarose gel to assess dsRNA integrity after soaking with

nematode J2s. After 16 hours of soaking, two thousand J2s were washed with DEPC-treated

water and snap frozen in liquid nitrogen for RNA extraction. For each dsRNA feeding

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treatment, ten tomato seedlings (cv. Grosse Lisse) were infected with 400 dsRNA-fed J2s to

assess infectivity.

5.3.6 Analysis of infection

After four weeks of infection, seven plants from each feeding treatment were carefully

uprooted, the roots washed gently, and observed under a dissecting microscope. The number of

galls present on the roots was counted for each treatment. To stain egg masses, roots were

treated with phloxine B by dipping the roots in the solution for five minutes after which they

were washed by dipping in clean water three to four times. Roots were pat-dried with a paper

towel and egg masses counted using a dissecting microscope. Root systems were then separated

from the stem at the collar, dried at 55 °C for 18 hours and weighed. The level of infection was

expressed as the number of galls and number of egg masses per gram of dry root weight.

After a further growth for three weeks, the remaining three plants for each treatment were

uprooted and the roots washed thoroughly. Adult female nematodes were dissected out of the

roots and stained with acid fuschin as described in Section 4.3.9. Adult females from plants

infected with treated nematodes were observed under the microscope (Olympus BX-51) under

bright field, and their appearance compared with those of controls.

5.3.7 Quantification of gene knockdown

After RNA extraction and DNaseI treatment, total RNA from dsRNA-fed nematodes was

purified and quantified using the Nanodrop. From each treatment, 500 ng RNA was used to

synthesise cDNA using the High capacity cDNA synthesis kit (Applied Biosystems) following

the manufacturer’s protocol.

One microlitre of this cDNA was used to quantify gene expression using qPCR assays. M.

incognita Actin (Accession no. BE225475) was used as internal control, while for the gene dcr-

1, qPCR primers designed from the region of the gene not included in any of the dsRNAs fed to

nematodes were used for all the seven dcr-1 targeting dsRNAs (Table 5.2). Transcript

abundance for three other RNAi effectors i.e. drsh-1, alg-1 and mut-2 was also tested. Primers

used are presented in Table 5.2. GoTaq® qPCR 2x master mix (Promega Corporation,

Australia) was used for all reactions in a Corbett RotorGene Quantitative Thermal Cycler

(Qiagen Pty Ltd., Australia) with five picomols each of the forward and reverse primers.

Cycling conditions were 95 °C for 5 min, followed by 40 cycles of 95 °C for 15 s and 55 °C for

60 s. All reactions were done in triplicates.

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Table 5.2: Primers used for quantitative RT-PCR assays. Amplicon lengths are indicated.

Relative gene expression was determined using the ∆∆CT method. Mean Ct value and standard

deviation was calculated using Rotor-Gene Q Series Software 1.7. Normalised CT value (∆CT)

relative to endogenous gene expression was calculated as described in Livak and Schmittgen

2001.

ΔCT of treatment= (Mean CT of target – Mean CT of Actin)

ΔCT of control= (Mean CT of target – Mean CT of Actin)

The calibrated value (ΔΔCT) was calculated by

ΔΔCT = (ΔCT of treatment - ΔCT of control)

Fold change in transcript expression relative to endogenous control was calculated as

Fold change = 2-(ΔΔC

T)

Relative change in gene expression was presented as fold change calculated with baseline

expression as 1.

5.3.8 Statistical analysis

The statistical software SPSSv20 (IBM Corporation, US) was used for data analysis. Nematode

infection and reproduction data was analysed using ANOVA and Tukey’s test for significance

analysis and comparison of means (p<0.05). For statistical analysis of qPCR data, Kruskal-

Wallis one-way analysis of variance was done to test significance at p<0.05 and 95%

confidence interval. Microsoft Excel Analysis ToolPak was used for construction of bar charts.

ID Primer 5′-3′ Amplicon

(bp)

qMiActin-F

qMiActin-R

TTGATGTAACAACCTCTGGGA

AATTCAAGGTTGAATTACTTGATCG

125

qMiDcr1-F

qMiDcr1-R

TCGTCGGGTGTTTGTGAATTA

ACATCCTCTTTCTGCAACTCTT

147

qMiDrsh1-F

qMiDrsh1-R

CAAGTGAATATCTTTACAAACAATTTCC

CCT GTG GAATAACCAAATATTTAACC

140

qMiAlg1-F

qMiAlg1-R

GGAATGCCAATTCAAGGTCAA

GCCAGGAAGTACAACACAAAC

129

qMiMut2-F

qMiMut2-R

CGACGATTGGCTTGCATTATT

GTTGATTGGCGTGTTCGTTTA

84

107

5.4 Results

5.4.1 Expression pattern of dcr-1 in J2 M. incognita

Both the antisense probe and sense control were used for DIG labelling of M. incognita J2s in

separate reactions. No signal was detected for nematode sections hybridised with the sense

probe. For the antisense labelled probe, the localisation of transcript appeared to be

predominantly in the intestine. There also appeared to be some expression along the nematode

body wall (Figure 5.1).

5.4.2 RNAi phenotypes after dsRNA soaking

J2 nematodes were soaked in solutions containing the seven dsRNA (dsD1 to dsD7), and with

dsgfp and without dsRNA for 16 hours. After soaking, 20 µL (20 µg dsRNA) of solution was

run on a gel to assess the integrity of the dsRNA after nematode soaking/feeding activity. No

apparent degradation of dsRNA was observed in any of the soaking solutions as sharp bands

were present for all dsRNAs visualised (Figure 5.2A).

When nematodes were studied under the microscope after 16 hours soaking, control J2s (no

dsRNA and dsgfp) were active and moved normally. The pattern of FITC fluorescence indicated

that the external solution had been taken up by the nematodes, and had spread throughout the

body (Figure 5.2B).

In contrast, J2s soaked in dsRNA targeting dcr-1 were inactive and straight, with slow

movement in the head region, whilst the lower body appeared paralysed. This phenotype was

consistent for each of the nematode dsRNA treatments targeting the 7 dcr-1 domains (Figure

5.2F).

(C)

BW

BW

BH

WB

BW

BW

(B)

I

(A)

Figure 5.1: Expression pattern of dcr-1 in M. incognita J2 nematodes as indicated by in situ

hybridisation. (A): Sense probe hybridisation shows no staining. (B): Staining observed in the

intestine (I). (C): Nematode body wall (BW) stained after antisense hybridisation. Scale bar =

50 µm.

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5.4.3 Nematode infection after dcr-1 RNAi

After 16 hours soaking in dsRNA, nematodes were then used to infect tomato plants. Four

weeks after infection, galls present on the roots were counted to assess the effects of RNAi on

host parasitism. No apparent differences were observed in gall morphology between controls

(soaked with no dsRNA or with dsgfp), and dcr-1 dsRNA-fed nematodes. There was also no

significant (p<0.05) difference between the number of galls/g of dry root for plants infected

with nematodes given control treatments. However, there were significant differences in the

number of galls/g dry root for four of the dcr-1 targeting treatments when compared to the

controls. These were for dsD1, dsD2, dsD3 and dsD7. The other three silencing triggers (dsD4,

dsD5 and dsD6) did not result in a significant reduction in the number of galls/g dry root on

tomato plants (Figure 5.3).

(A) M 1 2 3 4 5 6 7 8 9

dsRNA

FITC

200 µm

(B)

S FITC

S

(D) (E) (F)

No

dsRNA

dsgfp dsdcr-1

(C)

Figure 5.2 (A): Feeding solution run on gel showing that the dsRNA was not degraded after

incubation with nematodes for 16 hours. M = 100bp DNA ladder; Lanes 1 to 7: dsD1 to dsD7,

Lane 8: dsgfp, Lane 9: No dsRNA. (B and C): FITC uptake by nematodes after soaking for

16 hours. Fluorescence is visible in the gut and at the stylet (S). J2s after 16 hours soaking

without dsRNA (D), with dsgfp (E). (F): Representative phenotype of J2s after 16 hours

soaking with any of the seven dsRNAs targeting M. incognita dcr-1. The one depicted in the

figure is after feeding the domain Dicer dimer (dsD3).

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Figure 5.3: Average (Mean ± Standard error) number of galls produced per gram dry weight of

tomato roots by the nematodes soaked without dsRNA, with dsgfp and the seven dcr-1 targeting

dsRNAs. Significance with respect to the no dsRNA control is indicated by * (p<0.05).

The largest reduction in the number of galls/g dry root was recorded for the dsRNA treatment

targeting the 5′ terminal region of dcr-1 mRNA (dsD1). It reduced infection by up to 61%.

DsD2 reduced infection by 37% while dsD3 affected nematode infection by 35%. The dsRNA

dsD7 targeting the region closer to the 3′ terminal end of mRNA again reduced infection

significantly, by up to 46%. Therefore, out of the 7 dsRNA triggers, four were able to reduce

parasitic success significantly and reduced nematode establishment on plant roots.

5.4.4 Differential effects of dsRNA to different target region on reproduction

To assess the effects of dcr-1 RNAi on nematode reproduction, the number of egg masses was

counted for individual plant roots after staining with phloxine B, and egg masses per gram dry

root weight were calculated. There was no significant difference between the egg masses

produced by nematode treatments without dsRNA and with dsgfp. When dsRNA to the three

mRNA regions at the 5′ end of dcr-1 transcript (i.e. by dsD1, dsD2 and dsD3), were fed to

nematodes, reproduction was affected significantly, with the highest impact delivered by dsD1

where 67% reduction was observed in egg mass production (Figure 5.4). Egg mass production

in nematodes treated with dsD2 and dsD3 was reduced by 48% (dsD2) and 56% (dsD3)

respectively (Figure 5.4).

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Figure 5.4: Average (Mean ± Standard error) number of egg masses produced per gram dry

weight of tomato roots by nematodes treated with no dsRNA, with dsgfp and with the seven

dcr-1 targeting dsRNAs. Significance (p<0.05) with respect to no dsRNA control is indicated by

*.

The nematodes fed with dsD4, dsD5 and dsD6 did not show any significant change in egg mass

production (p<0.05). Egg mass production was reduced for nematodes that were soaked with

dsD7, but this reduction was not statistically significant (p<0.05) when compared to controls (no

dsRNA and dsgfp).

5.4.5 Nematode development is affected by dcr-1 RNAi

After seven weeks infection of tomato roots, adult female nematodes were dissected from the

roots, stained with acid fuschin, and observed under the microscope. Although the same gene

was targeted for all treatments, there were differences in morphology of the mature females

developed from J2s treated with dsRNAs corresponding to the different segments of the dcr-1

mRNA. Nematodes soaked with dsD1, dsD2, dsD3, dsD6 and dsD7 developed into smaller size

females compared to the controls (Figure 5.4).

This phenotype was very obvious for females developed from J2s fed on dsD1, dsD2, and dsD6.

The percentage of mature females with the characteristic smaller size was 53% for dsD1, 57%

for dsD2 and 51% for dsD6. About 5% of each of the females developed from J2s previously

fed dsD1, dsD2, dsD4, dsD6 and dsD7 were transparent to varying degrees.

111

5.4.6 Target quantification

Transcript abundance of dcr-1 in J2s was assessed after 16 hours soaking in dsRNA (Figure

5.6). Expression of dcr-1 in no dsRNA-treated nematodes was taken as the baseline for analysis.

For analysis of variance, the assumption for Levenes Statistics was not met for the data so the

non parametric Kruskal-Wallis test was done to analyse significant differences between

treatments. Significant differences were found between transcript abundance for the seven dcr-1

dsRNA and dsgfp treated nematodes. The three targets at the 5′ end of dcr-1 mRNA resulted in

down regulation of gene expression although it was variable for the different targets. Highest

decrease in transcript level was observed for the dsRNA (dsD1) targeting the first domain

coding region (DEAD-like Helicase).

DsD5 and dsD7 treatments showed the largest increase in expression; followed by the similar

up-regulation resulting from soaking with dsD4 and dsD6. With the exception of dsD5, up-

No dsRNA dsgfp dsD1

dsD2 dsD3 dsD4

dsD5 dsD7 dsD6

Figure 5.5: Adult M. incognita females dissected from tomato roots. Nematode (J2)

treatments were: no dsRNA, dsgfp and seven dsRNAs (dsD1 to dsD7) targeting the dcr-1

gene before infection. Females were stained with acid fuschin. Scale bar = 500 µm.

112

regulation of the dcr-1 transcript resulting from soaking increased for dsRNA corresponding to

sequences distal to the 5′ end of the gene. Surprisingly, dsgfp ingestion also resulted in up-

regulation of dcr-1 transcript in nematodes after 16 hours.

Figure 5.6: Relative quantification of transcript abundance for nematodes treated with dsgfp

and dsRNA targeting different regions of M. incognita dcr-1 transcript. Gene expression for the

no dsRNA control was used for data normalisation, and is represented by the baseline value of

zero. All treatments were significantly different from the no dsRNA treatment (p<0.05).

5.4.7 RNAi of dcr-1 affects other RNAi effectors

To investigate how changes in dcr-1 expression affected expression of other RNAi effectors, the

expression of three other RNAi effectors (drsh-1, alg-1 and mut-2) was quantified in dcr-1

targeted nematodes, and those fed with dsgfp. For this experiment, nematodes fed with dsD1

and dsD5 were chosen because they represented the extremes of expression of dcr-1 treatments.

Expression of these four genes in nematodes fed with dsgfp were also assessed. Expression in

control treatment with no dsRNA was used to normalise expression of the target genes.

Relative transcript abundance is presented as fold change in gene expression (Figure 5.7). Data

was normalised against actin as an endogenous transcript control. The results show that dcr-1

transcript expression was up-regulated when dsgfp was fed to nematodes. The other three

components of the pathway also showed change in transcript abundance i.e. increase in drsh-1

but decrease in expression of both alg-1 and mut-2. However, the two fragments targeting the

dcr-1 transcript at different regions generated different transcript levels for the RNAi effectors.

The transcript expression of dcr-1 in response to dsD1 was 53% less than the control whereas

for dsD5 it increased by 2 folds after 16 hours treatment.

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The transcripts for drsh-1, alg-1 and mut-2 were also affected by the two triggers. With the

down-regulation of dcr-1 in response to dsD1, drsh-1 expression increased by 27% (0.27 folds)

of the normal level while in response to dsD5 this gene was down-regulated by 56% (Figure

5.7B,C).

The genes alg-1 and mut-2 were down-regulated in response to dicer targeting triggers more

than in response to dsgfp feeding. Interestingly, drsh-1 expression showed an inverse

relationship with dcr-1 expression when dcr-1 was being targeted by dsRNA (i.e. drsh-1

Figure 5.7: Relative gene expression of three RNAi effector genes after nematodes were

treated with dsgfp and dsdcr-1. The zero or baseline indicates expression in the no dsRNA

treatment. (A): Expression level of dcr-1, drsh-1, alg-1 and mut-2 in dsgfp fed nematodes.

(B and C): Relative expression pattern of dcr-1 and other RNAi pathway genes (drsh-1, alg-

1 and mut-2) in nematodes fed with dsD1 and dsD5 respectively.

114

upresulated when dcr-1 down-regulated and vice versa) but was up-regulated when dcr-1 was

upregulated in response to dsgfp feeding (Figure 5.7).

5.4.8 Target and trigger properties affecting RNAi efficiency

The dsRNA targeted to 5′ end of the target mRNA produced the greatest effect on nematode

viability and reproduction. To further understand the different results from targeting the same

gene with different triggers, the seven dsRNA triggers were analysed for their GC percentage.

The GC percentage of the dsRNAs used was between 30.9% and 37.7%, with dsRNA lengths in

the range of 125-272 bp (Figure 5.8A). No relationship could be established between trigger

GC% and reduction in infection. Reduction in infection was in agreement with transcript

reduction for the three triggers near the 5′ of the target mRNA. To examine if the RNAi

efficiency was affected by the secondary structure of target mRNA, secondary structure of the

Midcr-1 mRNA (predicted from the contig CABB01000157 in Chapter 2) was constructed

using the RNAfold Web server (http://rna.tbi.univie.ac.at/cgi-bin/RNAfold.cgi). The predicted

structure for the dsRNA targeted regions is provided in Figure 5.8B.

(A)

(B)

Figure 5.8 (A): GC percentage and length of dsRNA targeting the seven regions of M.

incognita dcr-1 gene. (B): Secondary structure of dcr-1 mRNA predicted by the program

RNAfold. The seven target locations are indicated with respective dsRNAs.

115

It was not possible to correlate the secondary structure of the mRNA at regions used for dsRNA

synthesis to the effects of RNAi they induced, although a low positional entropy and higher

base-pair probability was calculated by the program for the region upstream of dsD4 (data not

shown). Interestingly, predicted secondary structures of the regions for dsD1 and dsD7 both of

which significantly affected nematode infection ability but showed opposite transcript

abundance, were in close proximity to each other (Figure 5.8B).

5.5 Discussion

In this study, dsRNA treatments designed to reduce expression of the dcr-1 gene did effectively

reduce infection levels and reproductive success of M. incognita. However, the extent of the

response also depended on the target region used for dsRNA in the soaking treatment. The three

triggers near the 5′ end of the dcr-1 transcript significantly reduced gene expression, nematode

infection, reproduction and development. This result is in agreement with the data published by

Arguel et al. (2012), who also found differential effects on target expression to dsRNA

treatments of the Calreticulin gene in response to different triggers in M. incognita. The GC% of

the regions targeted was similar; therefore other mechanistic or thermodynamic properties of

target RNA, such as accessibility and disruption energy, may be involved. Target accessibility

has been reported to play a role on efficiency of RNAi and contribute to its efficiency by up to

40% (Shao et al. 2007). However, the length of the dsRNA sequence used did appear important,

since longer dsRNA sequences seemed to have a lower effect on infection.

Interestingly, the trend of transcript change seems to correlate with the reduced infection and

reproduction of nematodes. This data also correlated with the quantitative gene expression data

for which gene silencing was observed for the same three targets for which the most reduction

in infection was observed. For the trigger dsD7, nematode infection reduced significantly but

transcript down-regulation was not observed. although up-regulated at the time of quantification

of transcript but possibly it did reduced later as reported for M. incognita genes where highest

silencing was observed 20 hours after washing nematodes off a four hour dsRNA treatment for

the gene Mi-crt while highest silencing was achieved after 44 hours for the gene Mi-pg-1 (Rosso

et al. 2005). The dcr-1 mutants in C. elegans have been reported as sterile and development

defective (Grishok et al. 2001). Developmental defects were observed for the parasitic

nematode M. incognita in this experiment too.

A clear finding of this study is that the RNAi pathway is activated when dsRNA enters the cells

as the dcr-1 mRNA was up-regulated along with drsh-1 while alg-1 and mut-2 were down-

regulated. The expression of these three seems to be affected by RNAi of dicer probably

because they play a role in the gene regulation cascade in an attempt to return to the optimum

116

balance of transcripts for the pathway. A complete gene expression profile for RNAi effectors

will probably contribute to understanding the role of each in this mechanism.

From previous studies, it seems that RNAi processes can exhibit a bi-phasic effect i.e. initial up-

regulation followed by down-regulation (Rosse et al. 2005; Arguel et al. 2012). However, in

this experiment the time of gene quantification (after 16 hours) showed both down-regulation

and up-regulation depending on the region of the gene targeted. The developmental defects

observed for dsD6 and dsD7 and infection reduction for dsD7 targeted nematodes possibly

resulted in response to down-regulation in gene expression at a later stage which then affected

nematode infection, reproduction and development.

In conclusion, RNAi of M. incognita dcr-1 affects nematode viability, parasitism and

reproduction. The role of the dicer in the RNAi cascade is very important and disturbing its

stability results in imbalance in the mechanism and possibly the activation of the miRNA

pathway to counteract the imbalance through altered gene regulation.

117

Chapter 6

General Discussion

118

6.1 Overview

In the work presented in this thesis, the RNAi pathway of the root-knot nematode M. incognita

has been studied in detail, together with that of other parasitic nematodes, with a focus on PPNs.

The overall aim of these studies was to determine the potential for genes in the RNAi pathways

as possible targets for nematode control. The knowledge generated could also be applied more

broadly to control other PPNs. Experiments were conducted to meet the following specific

research objectives (Chapter 1), which were

1. To apply comparative bioinformatics and molecular tools, and information available for

C. elegans and animal parasitic nematodes, to genes involved in the RNAi and miRNA

pathways of RKNs, combined with using ESTs and genomic data available for M. hapla

and M. incognita.

2. To study the effect of down-regulating expression of the identified RNAi

genes/effectors via in vitro feeding of dsRNA on the survival and/or parasitism of M.

incognita.

3. To investigate the effect of using dsRNA corresponding to different parts of an RNAi

pathway gene for down-regulation of expression and assess the effects on nematode

parasitism and reproduction and relative expression of other RNAi pathway

components.

4. To investigate the possibility of controlling M. incognita via host-induced

siRNA/dsRNA of RNAi effectors.

All these specific aims have been addressed successfully, with the generation of new

information for each of the aims.

In summary, considerable differences were found in the repertoire of RNAi pathway

components present in parasitic nematodes compared to those found in the free-living model

nematode C. elegans. There were also differences in domain organisation of some genes.

Twenty seven effectors of RNAi were cloned and targeted through in vitro feeding of dsRNA to

J2s of M. incognita and their infection and development was assessed. The results showed that

there was up to 90% reduction in infection after targeting the dcr-1 gene. The effects of

silencing other components of the RNAi pathways also resulted in reduced nematode

119

reproduction, depending on the target gene and other factors. One observation was that RNAi of

components of the miRNA pathway resulted in significantly impaired nematode development.

Transgenic plants expressing dsRNA to a subset of target genes identified from in vitro soaking

experiments had reduced nematode infection, with a reduction in infection of up to 89%. All the

transformed plants expressing target gene dsRNA exhibited reduced nematode infection by 50%

or more (based on 31 events of 7 genes). Adult female nematodes that did develop on transgenic

plants showed abnormal development, and the results obtained provide some priority gene

targets for further progression for nematode control.

DsRNA targeted to different regions of the dcr-1 gene affected J2s to different extents, with

triggers closer to the 5′ end of the gene being more effective as evidenced by reduced transcript

level, fewer nematodes infecting roots and reproducing. In further studies, it was also found that

in vitro RNAi of dcr-1 also affected expression of other genes in the RNAi pathway, suggesting

that the expression of several other non-targeted genes could be affected, and so by targeting

expression of a single gene more profound effects on nematode viability may result. Some of

these aspects are discussed in more detail below.

6.2 Effectors of small RNA pathways of RKN

The first objective of this project was to identify genes involved in the RNAi pathway of RKNs

using ESTs and genomic data available for M. hapla and M. incognita. The availability of

whole genome sequences of M. incognita and M. hapla made it possible to confirm domain

architecture of genes compared to C. elegans and to those of the animal parasitic nematodes A.

suum and B. malayi. Apart from the identification of genes found in these nematodes, this study

also showed that there were some disparities in comparison to previously published literature on

this subject (Rosso et al. 2009; Dalzell et al. 2011). Such differences may be partially attributed

to the stringency used and differences between the various data analysis software packages used

in the analyses.

The two Meloidogyne spp. seem to lack genes present in the RNAi pathway of C. elegans, these

relate to RNAi spreading, inhibitors, argonautes and genes involved in heritable RNAi.

Differences in functional domains were also found between species. Despite these differences,

the considerable body of literature now published on RNAi in relation to root-knot nematodes

provides clear evidence of a robust functional RNAi mechanism in these species, backed by in

vitro feeding data or from HIGS (reviewed in Lilley et al. 2012). In C. elegans, RNAi can

persist over several generations, through inheritance of nrde-3 associated siRNAs and histone 3

lysine 9 methylation marks on the target gene locus, while heritable transcriptional silencing has

120

been reported in the absence of trigger, which depended on the target gene used (Burton et al.

2011; Vastenhouw et al. 2006). Although the heritability of RNAi has not been investigated in

parasitic nematodes, there is some evidence for persistent down-regulation of an in vitro

targeted gene over generations for M. incognita (Gleason et al. 2008). This down-regulation

aided in plant infection and may be an epigenetic effect as a result of selection pressure and not

inherited RNAi as reported for C. elegans. This indicates that functional studies should be done

to explore the possibility that RNAi in PPNs might be heritable, and should be combined with

study of the functions of the genes drh-1, eri-1, tsn-1, which show differences in domain coding

regions with C. elegans orthologs.

6.3 In vitro RNAi as a functional analysis tool for parasitic nematodes

RNAi in C. elegans has predominantly been used as a tool to study functional genomics, but for

parasitic nematodes it has been used more to probe parasitism genes to increase understanding

of nematode-plant interactions, and to investigate the potential to use RNAi as a strategy for

nematode control. The second objective of this research was to study the effect of down-

regulating expression of genes in the RNAi pathways identified in the first part of the work, via

soaking with dsRNA, on survival and/or parasitism of M. incognita.

The results obtained in general are in agreement with similar studies on target knock-down and

reduced infection levels after dsRNA soaking. The observation that there can be an initial

increase in transcript level has also been observed after RNAi in PPNs (Rosso et al. 2005;

Bakhetia et al. 2008). This observation suggests that nematode metabolism responds to

perturbation of individual components of a pathway, and that there is active gene regulation

which responds to altered transcript levels. Up-regulation of the targeted transcript was

observed for 9 out of 16 tested genes in this study, and this suggests that down-regulation of

components of the RNAi pathways is not straight forward, and that to some extent, expression

levels compensate for perturbations in the pathway. It may be that higher trigger concentrations

might be more effective in knocking down genes for which up-regulation was found after

treatment, but responses to RNAi do not necessarily improve with increase in trigger

concentration: in some cases lower concentrations of dsRNA in soaking solutions are more

effective than higher concentrations (Sukno et al. 2007).

One issue to consider is that in vitro RNAi is a transient process, and may not reflect the longer

term effects of down-regulating a particular gene on a continuous basis. Nevertheless, it is a

valuable tool to provide an initial assessment of the effects of gene knockdown of potential

target genes in PPNs. In support of this argument, RNAi of a high proportion of the genes

studied did result in reduced infection and in developmental defects of adult females, indicating

121

that soaking in dsRNA can induce long lasting effects. Added to the factors considered, there

are a range of variables in the experimental conditions used in soaking experiments, such as the

optimum time of soaking, when down–regulation of a target gene actually occurs after dsRNA

ingestion, the target gene itself and the regions selected to generate dsRNA. Nevertheless, in

vitro RNAi screening of a larger number of target genes does enable the screening out of the

best prospective candidate genes from pathways studied. From this screen a subset of priority

targets were chosen, and progressed to study their effects on nematode infection and

reproduction with delivery of dsRNA via transgenic plants, i.e. by HIGS.

6.4 HIGS for RKN control

To investigate the possibility of controlling M. incognita damage via HIGS using RNAi

pathway effectors/genes as targets, transgenic A. thaliana plants were generated expressing

hpRNA to seven genes selected from the in vitro screen of 27 initial genes identified in the

pathway. Although there was some variability in the results, nematode infection was

significantly lower in plants expressing dsRNA than in control plants for events of all seven

genes tested. The greatest reduction in infection of 89% was for the target gene drsh-1.

In considering why reduction in infection after in vitro RNAi of a target gene does not

necessarily translate quantitatively for host-delivered RNAi, the most obvious explanation is

that the amount of trigger ingested by the nematodes in the two situations differs both

quantitatively and qualitatively. In soaking experiments, 1 µg/µL of dsRNA is present in the

external solution, whereas in host-delivered dsRNA it is not clear how much dsRNA is ingested

by the feeding nematode. In addition, the dsRNA in plant cells is likely to have been pre-

processed by the plant’s RNAi machinery, to generate the spectrum of siRNAs that are normally

present in plant cells. Higher concentrations of dsRNA provided in vitro sometimes increases

the gene silencing effect, but in other cases it does not. Perhaps higher concentration of dsRNA

can overwhelm the nematode RNAi pathway, but they can sometimes recover after the soaking

period. It is probable that transgenic plants deliver a lower concentration of dsRNA, over a

much longer time period, but in the form of pre-processed siRNAs. A better understanding of

the events that occur in transgenic plants requires further study to analyse the siRNA levels in

them, and whether the siRNAs that a nematode would generate itself from long dsRNAs differ

from those processed by the host plant. The HIGS studies undertaken in this research were done

in a growth chamber under controlled conditions. Host-delivered RNAi of the same gene might

act differently in field conditions, where plants may combat multiple stresses simultaneously.

Both short-term RNAi and long-term RNAi affected nematode development considerably,

indicating lack of complete recovery of nematodes from the effects of RNAi.

122

The fact that a 400 bp hpRNA corresponding to gfp expressed via a root specific promoter

silenced GFP expression in shoots of Arabidopsis plants (Liang et al. 2012) suggests that long

dsRNAs are processed to siRNAs, and that the silencing signals can be transported around the

plant. However, in the absence of a target message sequence in plant cells there is no

amplification of the signal.

The transgenic plants challenged with nematodes were heterozygous because of time

constraints. A better measure of the extent of HIGS would have been obtained from the next

generation of transgenic plants, challenging only those that were homozygous for the transgene.

Nevertheless, this study indicates that clear results can be obtained using heterozygous plants,

since there were clear reductions in infection success and modified development of adult

females. The knowledge generated should now be applied and tested in commercial crop species

to further determine their potential to reduce nematode infection. In many cases crop plants

have been selected for some level of resistance to RKNs, and so additional resistance conferred

above that present in the most resistant genotypes available, could result in very effective

control. That a reduction in nematode infection of 89% was found for the target drsh-1 suggests

it would be a very good candidate to add to existing resistance in commercial genotypes, and

could result in effective control of M. incognita in commercial agriculture.

6.5 The target region of a gene affects RNAi effectiveness

The effectiveness of RNAi depends on factors such as sequence composition and target region

chosen. This aspect was studied using targets for dcr-1 gene, which has a series of distinct

functional domains. Differences in the reduction of gene expression and infection after using

different regions from this gene were evident. These did not correlate with GC content, and so

other factors must explain this result. One explanation is that not all target sequences chosen

were equally effective in generating siRNAs, and this resulted in dfferential transcript

abundances after RNAi treatments. A similar result has been reported, that is different transcript

abundance after using different sequences from the same gene (Sukno et al. 2007; Arguel et al.

2012). Whether this efficiency depends on target thermodynamics or trigger composition

requires further investigation through more in vitro screens of genes. Another feature is that

after dsRNA soaking down-regulation in the dcr-1 transcript was observed using qRT-PCR for

the target regions close to the 5′ end of mRNA. That is, the target sequence expression was

down-regulated after soaking for 16 hours for those regions but up-regulated for other target

regions. The regulation of each target gene chosen may be different, and so this kind of

observation indicates another level of complexity that needs to be understood before the most

effective silencing of target genes can be achieved.

123

In the initial feeding experiment for RNAi of dcr-1, infection was reduced by 90% (Chapter 3).

However, when more detailed studies were done on this target gene (Chapter 5), the level of

reduction of infection was lower. This is not a surprising result since there are many comments

in the literature that relate variable or inconsistent results for RNAi of PPNs. Explanations range

from experimental variables such as seasonal variation of plants or nematodes, as noted for both

the root lesion nematode P. vulnus and for M. incognita (Britton 2001; Arguel et al. 2012).

It should be noted that the experiments for this study were done in spring, which is the

beginning of active season for M. incognita, while the first dcr-1 RNAi experiments were done

during autumn when the nematodes are less active. It is therefore possible that during spring and

summer, nematodes are more resilient to RNAi and can recover more quickly from its effect.

Regardless of the differences in response, the treatments even though transient, disturbed the

gene regulation machinery enough to lower infection, reduce reproduction and cause defective

development.

6.6 Future directions

The construction of plant transformation vectors for all the 27 genes cloned is a useful resource

which can be used to facilitate future HIGS studies in plants after transformation to express

hpRNA. In addition, the transgenic Arabidopsis plants could be taken to subsequent

generations, and analysed further such that only plants with homozygous single copy inserts are

used to study responses to nematode challenge more precisely. The transgenic plants can also be

used to study transcription and processing of foreign RNA to quantify the amount of gene

silencing signals (long dsRNA/siRNAs) produced. This will add knowledge on the

amount/concentration of triggers delivered to the parasite, and which plant derived siRNAs are

most effective for silencing targets in nematodes.

A further question that can be addressed is the stability of RNAi based resistance in following

generations. Under selection pressure, PPNs can overcome natural resistance genes, for example

virulent resistance-breaking strains of H. glycines develop when highly resistant soybean

cultivars are grown (Zheng and Chen 2011). The question for RNAi-based resistance is whether

it will be durable in a field situation or not. Although there is evidence for long term expression

of an RNAi-based trait for resistance to potato virus Y (P. Waterhouse, personal

communication), a transgene can be methylated to reduce its expression, and a mutation that can

by-pass the control could also be selected for in nematode populations. These possibilities are

no different from what can happen for any transgenic trait or indeed after application of

chemical pesticides, and is not necessarily limited to RNAi-based traits. Nevertheless this aspect

needs further study.

124

In C. elegans long dsRNA is diced into ~22-23 bp siRNAs, while for M. incognita, synthetic 21

bp siRNAs have demonstrated to confer efficient gene knockdown (Arguel et al. 2012). In

contrast, the predominant siRNAs generated in plants is 21 bp, with 22 bp and 24 bp siRNAs

also produced. Thus there may be an imbalance in the sizes of siRNAs produced by plants on

processing long dsRNA, and the preferred product used for gene silencing in nematodes. The in

vivo siRNA dicing process of long dsRNAs in PPNs requires further investigation. However,

recently a possible solution to this question has become evident. That is, plastids in plants, being

derived from cyanobacteria, lack the ability to process long dsRNAs. In the report by Zhang et

al. (2015), who were studying control of Colorado potato beetle by gene silencing, these authors

claimed 100% control through providing long dsRNA via plants with transformed plastids. The

two factors here are that there are many copies of plastid DNA in each plastid, and many

plastids in each cell. This ingestion of cells with transformed plastids provides large amounts of

long dsRNA, which can then be processed by the pest machinery rather than the plant RNAi

machinery. A similar approach, targeting root plastids, could increase the effectiveness of host-

delivered RNAi against PPNs, although the possible filtering properties of feeding tubes formed

by root-knot and cyst nematodes might prevent ingestion of plastids.

Another new tool that has emerged is ‘genome editing’. It can be used as a tool for functional

genomics studies and to silence expression of a targeted gene in a plant, possibly without

insertion of any external sequence. Such plants should be regarded as mutants by regulators,

rather than as genetically modified (GM) plants (Fosu-Nyarko and Jones 2015). At present this

is limited to editing the plant genome, and cannot be used to edit a nematode genome via a

plant. If genome editing were used for nematode control, a converse situation needs to be

addressed, that is, discovery of plant susceptibility genes to nematode infection, which could be

silenced by genome editing, so conferring resistance.

For an RNAi approach for nematode resistance to be commercialised, there are a number of

policy and regulatory hurdles, followed by issues of public acceptance, and other aspects such

as patenting and the costs associated with implementation. These aspects are discussed in detail

by Fosu-Nyarko and Jones (2015).

6.7 Conclusions

Significant new knowledge was generated in the work presented here in relation to the RNAi

pathway of M. incognita and the reduction of infection of the RKN M. incognita achieved by

down-regulating genes involved in the RNAi pathway. When miRNA pathway genes were

targeted, nematode development was profoundly affected, and this suggests that target genes

taken from this pathway could provide new candidates for an RNAi based control strategy.

125

Measurement of changes in target gene expression after soaking experiments showed that RNAi

targeted genes were up-regulated after soaking, but when soaked nematodes were used to infect

plants, the expression might subsequently be down–regulated since nematode infection,

reproduction and development could be reduced.

Using seven priority gene targets identified from soaking experiments, host-delivered dsRNA

was investigated using transgenic Arabidopsis plants, which constitutively expressed hpRNA

targeting M. incognita dcr-1, drh-3, vig-1, mut-7, drsh-1, pash-1, rha-1 genes. The response to

nematode infection of these transgenic plants was one of reduced nematode infection and

reproduction. Although due to time constraints, the plants screened for nematode responses

were not all homozygous for the hpRNA transgenes, significant reduction in infection was

observed in this segregating generation for events of all the genes tested.

After soaking treatments of 16 hours, in some cases there was a clear phenotype (paralysis,

abnormal wavy movement etc), yet from 9 out of 16 genes tested there was up-regulation of

expression of the targeted gene. Maybe the soaking procedure resulted in overloading the RNAi

machinery of the J2s, or compensatory responses in transcription of pathway genes occurred

which also affected expression of down-stream genes. The period of 16 hours or more for

soaking is commonly used, but perhaps a shorter period of soaking might be more effective.

Overall, the screening of many target genes by soaking in dsRNA followed by plant infection is

a valid strategy to identify candidate genes most likely to be effective in conferring improved

resistance in transgenic plants expressing hpRNAs, although the correlation is not exact.

Differences may relate to the form of the dsRNA ingested by nematodes on soaking or from

transgenic plants.

Different responses to nematode infection found when using dsRNA from different regions of

the dcr-1 gene also demonstrates that the exact sequence of a target gene used is critical: not all

sequences generate the same intensity of siRNAs in plants or the same level of gene silencing.

A better understanding of the principles which underlie this phenomenon is needed to select the

best target sequences, i.e. those most likely to cause the strongest reduction in target gene

expression. The potential for development of improved resistance to RKNs via transgenic plants

is supported by the work undertaken in this thesis, but it also highlights aspects of the

mechanisms involved that are not yet fully understood.

126

APPENDIX Supplementary Table 2.1: Number of significantly mapped contigs for the four parasitic nematodes to C.

elegans genes with the contig IDs of the sequence coding for the respective effector protein.

C. elegans

Gene

No. of significant matches (Contig ID of gene coding sequence)

A.suum B. malayi M. hapla M. incognita

Spreading Proteins

sid-1 3 (ANBK01003199

AMPH01007595

AEUI02000078)

2 (AAQA01000131) - -

sid-3 100(ANBK01007000) 59(AAQA01000282) 26 (ABLG01000030) 30 (CABB01000892)

rsd-3 6 (AMPH01006102) 4 (CAPY01004512) 3 (ABLG01000350) 6 (CABB01006346)

xpo-1 6 (AEUI02000555) 2 (CAPY01003870) 1 (ABLG01001363)

3 (CABB01004119

CABB01002745)

xpo-2 6 (AMPH01015976

AEUI02000107)

3 (CAPY01001063) 1 (ABLG01000755) 1 (CABB01000462)

xpo-3 5 (AMPH01020037) 2 (CAPY01005588) - -

haf-6 90 (AMPH01011416) 30 (CAPY01002522) 14 (ABLG01001289) 17 (CABB01001990)

Dicer and Associated Genes

dcr-1 22 (ANBK01006853

AMPH01008524

AEUI02001038)

5 (CAPY01005536) 3 (ABLG01001138) 3 (CABB01000157)

drh-1 12 (AEUI02001000) 7 (AAQA01001229) 5 (ABLG01000164) 8 (CABB01006008)

drh-3 10 (AEUI02000457) 5 (AAQA01000086) 4 (ABLG01000567) 4 (CABB01002056

CABB01002184)

pir-1 13 (AMPH01008932) 6 (AAQA01002333) 2 (ABLG01002012) 4 (CABB01007043)

drsh-1 8 (AEUI02000028) 3 (AAQA01000005) 1 (ABLG01000521) 1 (CABB01000477)

pash-1 5 (AAQA01000310) 1 (ABLG01000935) 1 (CABB01004277)

RISC

ain-1 - 6 (AAQA01000363) - -

tsn-1 4 (ANBK01000604) 3 (AAQA01000346) 1 (ABLG01000878) 3 (CABB01000055)

vig-1 3 (AMPH01002339) 2 (AAQA01001685) 1 (ABLG01000254) 1 (CABB01000081)

RNAi Amplification

smg-2 5 (AMPH01007249) 6 (CAPY01005799) 3 (ABLG01000485) 15 (CABB01000694

CABB01008394)

smg-6 3 (AMPH01018798) 2 (AAQA01000011) 2 (ABLG01000285) 1 (CABB01000011)

ego-1 12 (AMPH01018474) 7 (CAPY01002544) 2 (ABLG01000591) 4 (CABB01000449)

rrf-1 AMPH01008866 CAPY01004404 ABLG01001448 CABB01000474

RNAi Suppressors

eri-1 3 (AMPH01004548) 5 (AAQA01000082) 1 (ABLG01000828) 2 (CABB01001883)

eri-5 8 (AMPH01019550) 1 (AAQA01007053) - -

eri-6/7 6 (ANBK01005918) 4 (AAQA01000760) 2 (ABLG01001054) -

rrf-3 10 (ANBK01005062) 7 (CAPY01003132) - -

gfl-1 8 (ANBK01002547) 4 (CAPY01002536) 1 (ABLG01000591) 1 (CABB01000795)

xrn-1 10 (AMPH01008093) 4 (CAPY01002459) - -

xrn-2 9 (AEUI02001203) 5 (AAQA01000030) 1 (ABLG01000657) 2 (CABB01001503

CABB01003205)

adr-1 3 (AMPH01003945) 4 (CAPY01001829) - -

adr-2 3 (AMPH01015223) 4 (AAQA01003201) - -

zfp-2 40 (ANBK01002640) 100

(AAQA01001012)

17 (ABLG01000194) 20 (CABB01002475)

127

C. elegans

Gene

No. of significant matches (Contig ID of gene coding sequence)

A.suum B. malayi M. hapla M. incognita

Nuclear RNAi effectors

mut-7 3 (AMPH01001240) 2 (CAPY01005167) 1 (ABLG01000582) CABB01000055

cid-1 7 (AMPH01013268) 2(AAQA01000021) 3 (ABLG01001487) 8 (CABB01002392)

ekl-4 4 (ANBK01004956) 4 (CAPY01005796) 1 (ABLG01000886) 1 (CABB01002952)

mes-2 7 (ANBK01002478) 4 (CAPY01005273) 2 (ABLG01001056) 4 (CABB01002321)

mes-6 3 (ANBK01006945) 2 (CAPY01003620) 1 (ABLG01000753) 1 (CABB010000967)

rha-1 42 (ANBK01001125) 25

(AAQA01000377)

9 (ABLG01000200) 20 (CABB01000079)

ekl-6 2 (ANBK01003842) 2 (CAPY01005521) - -

mut-2 16 (AMPH01013881) 4 (CAPY01003131) 4 (ABLG01000108) 10 (CABB01002462)

zfp-1 10 (ANBK01000381) 6

(AAQA01000140)

2 (ABLG01000109) 2 (CABB01000945)

Argonautes

PAZ,

PIWI and

DUF1785 (alg-1, alg-

2, alg-

4/tag-76,

T22B3.2,

T23D8.7)

ANBK01003974

AEUI02000475

AEUI02000675

CAPY01000537

CABB01000336

PAZ and

PIWI (R06C7.1,

F58G1.1,

rde-1,

C16C10.3,

ppw-

1/sago-2,

ppw-2,

sago-1,

csr-1,

T22H9.3,

ergo-1,

prg-1, prg-

2,

F55A12.1,

nrde-3,

Y49F6A.1,

C14B1.7)

AMPH01030278

AMPH01002307

ANBK01003950

AMPH01020161

AMPH01003096

AEUI02000283

CAPY01004823

AAQA01000069

ABLG01001935

ABLG01001452

ABLG01000197

ABLG01000685

ABLG01001306

CABB01002242

CABB01000355

CABB01001343

CABB01000488

CABB01002924

CABB01000184

CABB01000681

CABB01000079

CABB01000330

PIWI

(C04F12.1,

ZK218.8

and

ZK1248.7)

AEUI02000059

ANBK01006280

CAPY01002834

CAPY01005262

ABLG01002012

ABLG01001620

ABLG01000816

CABB01000426

128

Supplementary Table 2.2: Similarity of protein domains of the RNAi pathways genes of C.

elegans (Ce) to four parasitic nematodes M. incognita (Mi), M. hapla (Mh), A. suum (As) and B.

malayi (Bm) based on query coverage and total scores.

Gene (Ce) tblastx Domains % Similarity with Ce Total

Score(Coverage)

%Similarity with Ce Total

Score(Coverage)

Mi Mh As Bm

Transport Proteins

sid-1 Sid-1RNA channel - - 254(34)

254(34)

226(30)

158(28)

sid-2 No Domain - - - -

sid-3 Tyrosine kinase domain 724(69) 644(74) 805(67) 790(71)

SH3 - - - -

GTPase binding - - - -

sid-5 No Domain - - - -

rsd-2 Rsd-2 - - - -

rsd-3 ENTH-Epsin 292(79) 229(78) 402(94) 287(83)

rsd-6 Tudor - - - -

xpo-1 IBN-N

Xpo1 379(96) 384(96) 313(67) 241(72)

CRM1 667(79) 798(93) 862(93) 681(93)

xpo-2 IBN-N 43.9(47) 43(47)

CSE 297(65) 360(71) 545(92) 445(72)

CAS-CSE1 44.6(32) 42.8(21) 391(65) 456(79)

xpo-3 Xpo1 - - 133(53) 257(65)

haf-6 ABC membrane 107(39) 465(87) 490(98)

ATP binding domain 860(92) 863(90) 897(88) 388(78)

Dicer and associated genes rde-4 DSRM - - - -

pash-1 WW 144(81)- As* 144(81)-As* 337(100)-As*

DS RBD 197(77)-As* 215(80)-As* 601(87)-As*

drh-1 DEXDc 204(34) 190(50) 263(57) 189(62)

HELICc 234(75) 231(75) 182(71) 205(61)

RIG-I_C-RD - - - -

drh-3 DEXDc 140(62) 234(79) 224(89) 245(95)

HELICc 124(54) 98.7(54) 115(54) 154(54)

RIG-I_C-RD 473(83)-Mh* 51(69)-Mh* 53(53)- Mh*

drsh-1 RiboIIIa 238(100) 240(100) 236(100) 325(100)

RiboIIIb 305(92) 313(92) 181(81) 264(94)

DSRM 146(92) 196(61) 156(61) 148(61)

dcr-1 Hel ATP-B 256(80) 185(55) 374(97) 406(92)

Helic-CT 74.8(75) 73.9(67) 131(60) 126(62)

DS-RBF 216(90) 293(92) 293(92) 222(91)

PAZ 292(76) 286(70) 519(71) 526(82)

RiboC1 416(79) 392(82) 432(80) 419(79)

RiboC2 521(83) 474(88) 496(94) 487(98)

DSRM 171(100) 108(100) 112(100) 146(100)

pir-1 Dual specificity

phosphatase

190(84) 168(74) 218(86) 321(91)

129

Gene (Ce) tblastx Domains % Similarity with Ce

Total score(Coverage)

%Similarity of Ce

with APN Total

score(Coverage)

Mi Mh As Bm

RISC ain-1 M domain - - - 54.4(29)

ain-2 M domain - - - -

tsn-1 SNc1 488(85) 382(85) 337(91) 425(91)

SNc2 367(84) 491(84) 507(90) 423(84)

SNc3 292(79) 323(80) 320(77) 376(90)

SNc4 244(90) 280(90) 366(96) 268(93)

Tudor 64.3(51) 63.4(53) 246(97) 329(92)

SNc5 36.2(36) - - -

vig-1 HABP4_PAI-RBP1 65.4(19)-

Bm*

33.1(9)-

Bm*

114(38) 140(65)

RNAi Amplification smg-2 UPF1_Zn_

bind

554(97) 362(97) 374(95) 346(95)

P loop AAA_30 445(95) 525(96) 629(100) 578(98)

P loop AAA_12 758(100) 838(100) 903(98) 883(98)

smg-5 PIN Smg5 - - - -

smg-6 EST1 80.5 (64)-

Bm*

99.9(80)-

Bm*

136 (98) 112(82)

EST1 DNA Bind 103(28)-

Bm

80(29)-

Bm

70.7(19) 155(40)

PIN Smg6 64.5(50)-

Bm*

38.6(22)-

Bm*

58.2(31) 104(62)

ego-1 RdRp 1240(72) 1030(79) 1457(85) 814(38)

482(32)

rrf-1 RdRp 831(56) 1146(75) 1298(85) 1253(82)

rde-10 Maelstrom - - - -

rde-11 No Domain - - - -

RNAi Suppressors Eri-1 411(95) 280(87) 337(83) 327(83)

eri-3 No domain - - - -

eri-5 Tudor - - 118(61) 71.8(56)

eri-6/7 P loop AAA_11 - 33.6(17) 47.8(20) 116(37)

P loop AAA_12 - 144(57) 41.8(11) 157(37)

eri-9 Zc3h12a-like

Ribonuclease NYN

domain

- - - -

rrf-3 RdRp - - 1379(79) 1510(75)

gfl-1 Yeats 229(95) 226(93) 426(97) 526(99)

xrn-1 XRN-N - - 968(100) 44.1(16)

xrn-2 XRN-N 1709(47) 2565(77) 2709(83) 2285(75)

adr-1 DSRM1 - - -

DSRM2 - - 37.7(45) 67.7(46)

A. Deamin - - 125(42) 137(44)

adr-2 DSRM - - - -

A. Deamin - - 255(54) 119(39)

lin-15b THAP - - - -

zfp-2 Zn finger Domains 343(50) 515(63) 542(67) 457(48)

130

Gene (Ce) tblastx Domains % Similarity with Ce

Total score(Coverage)

%Similarity of Ce

with APN Total

score(Coverage)

Mi Mh As Bm

Nuclear RNAi Effectors mut-7 Mut-7 36.8(7) 109(19) 178(38) 172(43)

cid-1 PAP & PAPtutase 331(15) 358(11) 412(37) 448(19)

ekl-1 Tudor1 - - - -

Tudor2 - - - -

ekl-4 DMAP1 177(58) 112(46) 232(50) 342(48)

mes-2 COG2940 214(75) 216(75) 222(68) 375(65)

SET 177(91) 181(91) 131(83) 211(75)

mes-3 No Domain - - - -

mes-6 WD40 31.8(19) 100(24) 70.4(55) 239(38)

rha-1 DSRM1 125(97) 124(97) 102(100) 104(91)

DSRM2 38.6(35) 38.6(35) 66.9(75) 35(60)

DEAD 492(98) 442(95) 604(99) 424(99)

HelCT 402(93) 248(93) 416(97) 249(94)

HA2 162(95) 166(95) 194(92) 159(91)

OB NTP 254(97) 294(97) 245(98) 288(89)

ekl-6 DUF2435 - - 45.1(40) 48.7(38)

zfp-1 Zfp-1 domains 286(40) 266(41) 513(82)

645(80)

mut-2 PAP &

NT_PAP_TUTase

213(26) 115(15) 149(40)

181(20)

ekl-5 7TM-GPCR-Srn - - - -

mut-16 No Domain - - - -

rde-2 No Domain - - - -

nrde-1 No Domain - - - -

nrde-2 DUF1777 - - - -

Nrde-2 - - - -

nrde-4 No Domain - - - -

131

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Effect of Knockdown of Genes Involved in the RNAi Pathway ... · Effect of Knockdown of Genes Involved in the RNAi Pathway on Root-knot Nematodes This thesis is presented to Murdoch