Myzus Persicae)

Molecular aspects of Potato Virus Y transmission by

aphids (Myzus persicae)

Saman Bahrami Kamangar

A dissertation submitted in partial fulfillment of the requirements for the PhD degree

in Bioscience Engineering

Promoters:

Prof. dr. ir. Guy Smagghe, Department of Plants and Crops, Faculty of Bioscience Engineering, Ghent University

Dr. ir. Kris De Jonghe, - Plant Sciences Unit, Flanders Research institute for Agriculture, Fisheries and Food (ILVO)

Dr. ir. Nji Tizi Clauvis Taning, Department of Plants and Crops, Faculty of Bioscience Engineering, Ghent University

Dutch translation of the title: Moleculaire aspecten van aardappel virus Y transmissie door bladluizen (Myzus persicae)

Please refer to this thesis as follows: Bahrami Kamangar, S. (2021). Molecular aspects of Potato Virus Y transmission by aphids (Myzus persicae). PhD thesis. Ghent University, Belgium.

ISBN: 9789463574259

Ghent University: Rector: Prof. dr. ir. Rik Van de Walle

Faculty of Bioscience Engineering: Dean: Prof. dr. ir. Marc Van Meirvenne

Aknowledgment

During the preparation of this thesis, I received great support and assistance. First, I would like to thank the Agricultural Research Education and Extention Organization (AREEO) in Iran for funding a part of my study. I would like to extend my sincere thanks to my supervisors Prof. dr. ir. Guy Smagghe, Dr. ir. Kris De Jonghe and Dr. ir. Nji Tizi Clauvis Taning for their consistent encouragement, support and guidance over the running of this research. I also wish to thank the Jury members Prof. Dr. Daisy Vanrompay, Dr. Olivier Christiaens, Dr. Jochem Bonte, and Dr. Stephan Steyer for reviewing and improving this thesis. I am grateful to Steve Baeyen, Lab Manager at the Institute for Agricultural and Fisheries Research, for his technical support. My wife Touba and son Siamand deserve special thanks for their patience, spiritual and emotional support. Without their understanding and encouragement in the past few years, it would have been impossible for me to complete my doctoral research. I wish to thank my parents and sisters who supported and always gave me positive energy. I owe a great debt of gratitude to my brother Dr. Barzan Bahrami Kamangar for his encouragement and assistance when I was working on my PhD. I gratefully acknowledge the help of my friends Dr. Khosro Mehdi Khanlou, Dr. Asad Maroufi, Serkewt Safaei, Dr. Amir Sadeghi, Hemen Piri and Dr. Hossein Hosseini Moghaddam.

Saman Bahrami Kamangar

III

Contents List of abbreviations ...... 1 Doctoral project context and research questions ...... 5 Chapter 1: General introduction ...... 8 1.1. The potato plant as staple food ...... 9 1.2. Plant viruses ...... 10 1.2.1. Plant virus transmission ...... 15 Non-circulative non-persistent transmission (NCNP) ...... 23 Persistent circulative, non-propagative transmission (CNP) ...... 23 Non-circulative semi-persistent transmission (NCSP) ...... 24 Persistent circulative, propagative transmission (CPP) ...... 25 1.2.2. Plant virus symptoms ...... 25 1.2.3. Important potato viruses ...... 26 1.3. Potato virus Y (PVY) and its characteristics ...... 37 1.3.2. Genome organization and proteins ...... 38 1.3.3. Genetic diversity and strains ...... 40 1.3.4. Host range ...... 46 1.3.5. Transmission ...... 47 Strain specific aspects and virus plant sources ...... 48 Impact of the virus receiving host plants ...... 49 Aphid and transmission efficiency ...... 50 Winged versus wingless aphids ...... 54 Impact of the feeding behavior ...... 54 Environmental conditions ...... 56 Plant volatiles, color and nutrients ...... 58 Control measures; Insecticides, straw mulch and mineral oil ...... 59 Vector plant hosts before virus acquisition ...... 60 Starving, acquisition accession period (AAP), inoculation accession period (IAP) and virus retention ...... 61 Molecular aspects of PVY transmission by aphids ...... 63 1.4. Techniques to study virus transmission ...... 68 1.4.2. Electric penetration graph ...... 70 1.4.3. Electron microscopy (EM) and labeling ...... 71 1.4.4. Immunological methods ...... 71 1.4.5. Molecular methods (q(RT-)PCR and (RT-)PCR) ...... 72 1.4.7. RNAi (RNA interference) ...... 75

1.4.8. Statistical methods and calculation bottlenecks ...... 79 Chapter 2: Potato virus Y (PVY) strains in Belgian seed potatoes and first molecular detection of the N-Wi strain ...... 82 2.1. Introduction ...... 82 2.2. Materials and Methods ...... 84 2.2.1. PVY screening and collection of PVY isolates ...... 84 2.2.2. Strain determination ...... 85 2.3. Results ...... 87 2.3.1. Prevalence of PVY types in Belgium ...... 87 2.3.2. Strain determination ...... 89 2.3.3. Symptoms, strain and potato cultivar relation ...... 94 2.4. Discussion ...... 95 Chapter 3: Quantity and transmission efficacy of an isolate of the Potato virus Y-Wilga (PVY N-Wi) by aphid species reared on different host plants ...... 99 3.1. Introduction ...... 99 3.2. Material and Methods ...... 100 3.2.1. Aphids, plants and virus isolate ...... 100 3.2.2. RNA‑transcript synthesis ...... 101 3.2.3. PVYN−Wi transmission bioassays ...... 102 3.2.4. PVY N-Wi quantification in the aphid stylet and intact whole body ...... 103 3.2.5. Preparation of aphid and plant samples, and RNA extraction ...... 104 3.3. Results ...... 104 3.3.1. RNA transcript and standard ...... 104 3.3.2. Transmission of PVYN−Wi by aphids ...... 105 3.2.3. Quantification of PVYN−Wi in the aphid stylet and intact whole body ...... 107 3.3. Discussion ...... 108 Chapter 4: The cuticle protein MPCP2 is involved in Potato virus Y transmission in the green peach aphid Myzus persicae ...... 1111 4.1. Introduction ...... 111 4.2. Material and Methods ...... 115 4.2.1. Plants, aphids and virus ...... 115 4.2.2. Target gene selection and double‑stranded (ds) RNA synthesis ...... 115 4.2.3. Aphid dsRNA feeding bioassay and RNAi ...... 117 4.2.4. qPCR expression analysis of CuP genes after dsRNA feeding ...... 118 4.2.5. Virus transmission ...... 119 4.3. Results ...... 119

4.3.1. Successful gene knockdown by dsRNA feeding for two CuPs ...... 119 4.3.2. Knockdown of mpcp2 in M. persicae led to reduced virus transmission ...... 120 4.4. Discussion ...... 121 Chapter 5: General discussion and future perspectives ...... 126 5.1. Variability of PVY strains in Belgium ...... 126 5.2. PVY transmission efficiency by different aphid species ...... 130 5.3. The host plant on which an aphid develops affect its PVY transmission efficiency and PVY quantity ...... 131 5.5. Future recommended work ...... 137 References ...... 139 Keywords: ...... 164 Samenvatting ...... 165 Kernwoorden: ...... 167 Curriculum vitae ...... 168

List of abbreviations

+ssRNA Positive-sense single-stranded RNA

AAP Acquisition Access Period

AFP Acquisition Feeding Period

ANOVA Analysis of variance

BCMV Bean common mosaic virus

CaMV Cauliflower mosaic virus

CD Common duct cDNA Complimentary DNA

CLSM Confocal laser scanning microscopy

CMV Cucumber mosaic virus

CP Coat (capsid) protein

CPNP Circulative, persistent, non-propagative

CPP Circulative, persistence, propagative

CPR Cuticular protein family

CPR1 Cuticular protein 1

CuP Cuticular Proteins

CYVV Clover yellow vein virus

DAS-ELISA Double Antibody Sandwich- Enzyme-Linked Immunosorbent Assay

DNA Deoxyribonucleic acid dsRNA Double-stranded RNA

EPG Electric penetration graph

FC Food canal

GFP Green fluorescent protein

HC Helper component

HC-Pro Helper component proteinase

HIV Human immunodeficiency virus

HR Hypersensitive response

IAP Inoculation access period

ICTV International Committee on Taxonomy of Viruses

IgG Immunoglobulin G

JKI Julius Kühn Institute

IPM Integrated Pest Management

KITC (1 letter of Amino Acid codes) Lys-Ile-Thr-Cys

KLSC (1 letter of Amino Acid codes) Lys-Luc-Ser- Cys

KLTC (1 letter of Amino Acid codes) Lys- Leu-Thr- Cys

KVSC (1 letter of Amino Acid codes) Lys-Val- Ser- Cys

Mpcp Myzus persicae Cuticular Proteins

NCNP Non-circulative, non-persistent

NCSP Non-circulative, semi-persistent

NGS Next generation sequencing

NP Nucleoproteins

ORF Open reading frame

P2 CaMV transmission protein pC3 Nucleocapsid protein 3 RNA-binding viral protein of Rice stripe tenuivirus

PepMoV Pepper mottle virus

PepYMV Pepper severe mosaic virus

PepYMV Pepper yellow mosaic virus

PLRV Potato leaf roll virus

PMV Peanut mottle virus

PSV Peanut stripe virus

PTK (1 letter of Amino Acid codes) Pro-Thr-Lys

PTNRD Potato tuber necrotic ringspot diseases

PTV Peru tomato virus

PVA Potato virus A

PVV Potato virus V

PVX Potato virus X

PVY Potato virus Y

PYBV Potato yellow blotch virus

QITC (1 letter abbreviation of Amino Acid) Gln -Ile-Thr- Cys

RDP4 Recombination detection program (version 4)

RH Relative humidity

RITC (1 letter abbreviation of Amino Acid) Arg-Ile-Thr- Cys

RJ Recombinant junctions (breakpoints)

RNA Ribonucleic acid

RPS2 Ribosomal protein S2

RSV Rice stripe tenuivirus

RTTC (1 letter abbreviation of Amino Acid) Arg- Thr- Thr- Cys

SBMV Soybean mosaic virus

SPFMV Sweet potato feathery mottle virus

SOC Super Optimal Broth medium

TA Transmission activation

TEV Tobacco etch virus

UPGMA Unweighted pair group method with arithmetic mean

UV Ultra violet

VPg Viral protein genome-linked

WPMV Wild potato mosaic virus

Y2H Yeast-2-hybrid

YMV Yam mosaic virus

ZYMV Zucchini yellow mosaic virus

Doctoral project context and research questions

A majority of plant viruses are dependent on vectors for their transmission and survival. Vectors that transmit plant viruses include mites, protists, fungi, insects and nematodes. Among these vectors, insects are the most common, with aphids accounting for the transmission of about 50% of insect-vectored viruses. Although vector-virus interaction is known to play a key role in the spread and epidemiology of plant viruses, several knowledge gaps still exist about the detailed mechanisms involved in the vector-virus transmission relationship. Potato virus Y (PVY) is an RNA virus and 5th most damaging plant virus and causative agent of potato damage worldwide that infects a wide range of hosts but potato is the most important host

(Scholthof et al., 2011). PVY is transmited in a non-circulative non-persistent (NCNP) manner by aphids. NCNP transmission is one of the simplest (still specific) transmission types and at the same time it has complexities and unknowns.

Transmission and strain variability are important components of PVY epidemiology with some unknowns and obscurities that still need to be understood in details. The questions about these obscurities inspired us to take a step towards better understanding PVY transmission in the following 3 topics that could improve knowledge of PVY epidemiology and management:

 Chapter 2 Identification of PVY strains in Belgium.

 Chapter 3 Understanding the relationship between PVY quantity in the stylet of

aphid, rearing plants and PVY transmission efficiency.

 Chapter 4 Identification of PVY receptors in the aphid stylet.

PVY as an RNA virus is always changing and adapting to new environments and cultivars. This genetic diversity and new recombinants provide capacity to overcome

PVY resistance in potato cultivars, and this has increased significantly in recent years.

PVY strains induce a variety of symptoms in potato cultivars and the resistance is a cultivar-strain interaction. Due to lack of new information on the prevalence of PVY strains in Belgium, which plays a key role in the selection of resistant cultivars, breeding programs and the integrated management of the disease, it was important to update and collect new information on the diversity, occurrence and significance of prevalent strains. Chapter 2 is providing a survey on the presence of PVY strains in

Belgium that are the main material in followed chapters of this thesis.

NCNP transmission of PVY and the presence of different aphid species as vectors promote the transmission of PVY in the farm, but the transmission efficiency of these aphids varies depending on the conditions and aphid entities. Behavioral factors

(including food preference and host plant interaction) or the quantity of virus particles in the stylet (as the essential organ in PVY transmission) may be the most significant factors influencing transmission efficiency. In addition, limited studies indicate the inhibitory effects of various substances (also plant materials) on the transmission of

PVY, although the mechanism remains uncertain. The transmission efficiency of PVY aphids can be affected by plants materials, so inhibition of virus lodging in aphid stylet may be one of the reasons for this. Assuming that the substances in aphid host plants before the virus is acquired will affect the virus content of the stylet and reduce the transmission efficiency of the virus, the effect of the host plant on the transmission efficiency and the quantity of viruses in the stylet of vector and non-vector aphids was evaluated in another section of this dissertation. The impact of host plants for aphid rearing (prior to virus acquisition) on virus quantity and transmission efficiency were evaluated as a part of this dissertation that is discussed in chapter 3.

The sequence of PVY genes is well known and the role of encoded proteins by these genes in the plant and in virus transmission (HC and CP) are well known, however,

the biochemical details of receptors and behavior of aphid vectors in virus transmission are less known. The chemical properties of PVY receptor/s in aphids were unknown until recently. Indirect evidence from viruses with similar transmission mechanisms

(NCNP and NCSP), such as ZYMV in the Potyvirus genus and CuMV in the

Caulimovirus genus, indicated that the cuticle proteins (CuPs) in the distal part of the aphid stylet may be PVY receptors. Accurate detection of these receptors can be useful in PVY transmission prevention and management approaches.

RNAi has been used as an effective tool with a bright future (in managing pests and researching the biochemical cycles of living organisms) to recognize receptors involved in the transmission of viruses by silencing their genes. However, the diversity of cuticle proteins, the sequence similarity and their expression as gene families can impair the efficacy of this method. Chapter 4 is about the efforts of CuP genes silencing of most efficient vector of PVY (Myzus persicae) by means of dsRNA.

In the last chapter, the results of these studies and related recent findings in other laboratories were discussed to explain what we have added and what should be added to the knowledge of PVY in Belgium and around the world.

Chapter 1

General introduction

Chapter 1

General introduction

1.1. The potato plant as staple food

About 7,000 years ago, the Andes habitants in South America started gathering and experiencing potato tubers as a food. While the plants produce inedible true fruits, the tuber is, and produces enough energy-rich nutrients. The tubers also acted as a vegetative ‘seeds’ and the early humans in Andes discovered how to propagate the crop. They started to transport and store the tubers for feeding of the society and they learned to plant and grow potato gradually. Today’s potato (Solanum tuberosum) has been selected from S. brevicaule populations from the Andean region and nowadays has become one of the most important staple foods in the world (Navarre and Pavek,

2014).

Figure 1.1. Total production, yield and area harvested of potato in Belgium (2000-2018). https://www.fao.org/faostat/en/#data

Potato has been cropped on 65,000-98,000 ha (13-15% of total crop lands) in Belgium since 2000-2019. It is the second most produced commodity after the wheat, range from 3.4 to 4.4 million tons, which is about 0.7 to 0.9% of world production and put

Belgium in 22nd ranks in the world (Faostat, 2021). The potato crop has been affected by drought in rainfed systems, as was the case in Belgium in 2018, resulting in a 30% loss of the crop (Sawyer et al., 2019). The high edible energy and dry matter of potato

(per ha per day) (Fig. 1.1 and 1.2) makes it the biggest production value crop

(US$/ha/day) compared to the other main staple food crops (wheat and rice) (Roots, tubers, plantains and bananas in human nutrition - Nutritive value, 2021).

Figure 1.2. Comparison of harvested area and production of wheat and potato in Belgium (2018).

1.2. Plant viruses

Viruses as assembled genetic materials (RNA or DNA) in coat protein subunits the nucleoproteins (NP) and sometimes enveloped in lipoprotein membranes. They are non-living obligate parasites that start replication only in suitable host cells and use own genes to exploit the host cells and profit from ribosome and other protein production systems and structural materials present in the host cell, for its replication

and cell to cell movement (Hull, 2014). They have specialized and adapted to different types of host cells including plants. A wide diversity of plant viruses consisting of a variety of different shapes and sizes, as well as different types of genetic materials

(Fig. 1.3), cause important diseases in many crops, and result in major economic losses all over the world (Agrios, 2005). The economic impact of plant viruses on crops is higher than what it seems to be, because the symptoms could be confused with nutrient deficiencies, abiotic stress, herbicide and hormone damage or its vector damage in plants. (Scholthof, 2011; Hull, 2014).

The plant viruses can't enter host tissues by themselves to multiply and propagate and must be carried and facilitated to enter living cells and introduce a new plant infection.

Unlike for animal viruses, virus movement in the plants is restricted to plasmodesmata.

They can’t enter or exit from membrane protected cells directly, since they are bound by a strong cell wall, composed by pectin and cellulose (Benitez-Alfonso et al., 2010).

They rely on a third party to spread between same or to next generations of host plants by means of mechanical wounds of contaminated instruments, or feeding and/or parasitizing arthropods, nematodes, parasitic plants and fungi-like vectors

(Mukhopadhyay, 2011). Moreover, they infect plant progenies during sexual (true seeds and pollen) and vegetative reproduction, with the last pathway the most likely to transmit the viruses. A high rate of virus spread and transport over long distance by vegetative reproduction organs, can increase virus importance in the crops such as potato. A case study on potato viruses pointed out that infections with potato leaf roll virus (PLRV), annually cause about £30 - 50 million losses in UK alone (Hull, 2014).

Figure 1.3. Diversity in genetic material and morphology of plant virus.

Viruses are classified by International Committee on Taxonomy of Viruses (ICTV) and

currently include 4 realms, 9 kingdoms, 16 phyla, 2 subphyla, 36 classes, 55 orders,

8 suborders, 168 families, 103 subfamilies, 1421 genera, 68 subgenera, 6590 species of which about 2, 3, 7, 12, 16, 34 (+6 unassigned), 127 (+45 unassigned) and 1624 are plant virus realms, kingdoms, phyla, classes, orders, families, genera, and species respectively. In addition, to date, 33 viroids and 143 satellite viruses that cause virus- like symptoms or modify virus symptoms are classified as virus-associated organisms

(ICTV master species list 2019.v1, 2021). Recent studies based on high-throughput sequencing (HTS) technologies (next generation sequencing) are “discovering novel viruses” at an increased pace, because of the vast amount of data that are being generated in a short time frame, and because former studies mainly focused on eminent problems caused by symptomatic viruses resulting in latent or cryptic viruses being ignored (Massart et al., 2017; Adams et al., 2018; Maclot et al., 2020; Rubio et al., 2020). Some of these viruses in a co-evolution process together with the plant host have gained a stable condition as persistent cytoplasmic viruses. They do not transmit horizontally, do not move from cell to cell and only infect their hosts for many generations vertically. Some sequences of the genome of these viruses have integrated with genomes of the host plants and they could increase the plant abilities

(Roossinck 2012b; Schoelz and Stewart 2018). The most diverse plant virus families are those of the begomoviruses (Geminiviridae) and potyviruses (Potyviridae) respectively (Genus: Begomovirus, 2021; Genus: Potyvirus, 2021). The nucleic acid properties (RNA or DNA, double or single stranded, the sense and segment of the genome), morphology and structure of the virion, genome sequence homology, biological and serological properties are the most important characteristics used to discriminate different levels of virus taxa (King et al., 2012) (Fig 1.4).

Figure 1.4. Potyviridae phylogenetic tree. Pairwise unrooted neighbor-joining tree of complete polyprotein sequences of representative viruses within the family Potyviridae. The tree was produced in MEGA 7 Branches supported by >70% of 100 bootstrap replicates are indicated. (Adapted from: Potyviridae, 2021)

1.2.1. Plant virus transmission

The transmission of viruses takes place by sexual or vegetative propagation, mechanically and/or through vectors. Knowledge on the transmission modes are essential to grasp virus diversity and evolution, as well as a critical step in surveillance

(da Silva et al., 2020). Mechanical transmission takes place in different ways. Plant viruses move from infected plants to wounded healthy plants in a same generation and/or the same growing season (horizontal transmission), and this through direct contact with contaminated soil, water, tools and/or plant tissues (roots and foliage)

(Fageria et al., 2015). Survival and transmission of viruses in at least 7 genera have been detected in water of canals, lakes and even oceans and in soils including for carmoviruses, cucumoviruses, diathoviruses, tobamoviruses, necroviruses, potexviruses, tombusviruses and sobemoviruses, all among the most stable viruses to survive outside a host plant (Mehle and Ravnikar, 2012; Hull 2014; Mehle et al.,

2014). In addition, some of these plant viruses were tracked in processed food (e.g. pigmented sauces), humans and wild animal guts and feces, (Zhang et al., 2005;

Rosario et al., 2009; Li et al., 2010; Phan et al., 2011; Scheets et al., 2011; Roossinck,

2012a and b). On the other hand, some viruses are difficult, or even completely fail to be transmitted mechanically, especially those that are restricted to trachea (xylem or phloem), e.g. members of the Closteroviridae (Walkey, 1991).

Transmission to the next generation (vertical transmission) is possible by infected vegetative propagules (bulb, tuber, scion, grafts, rootstocks) and /or sexual (pollen and seed) propagation (Walkey, 1991). About one-seventh of all plant viruses are known to be transmissible through infected seeds, the basic input for most cultivated crops in agriculture, and are responsible for virus transmission over longer distances and time periods. In addition, they not only reduce seed germination, but also plant

vigor. Infected seed coats (e.g. for tobamoviruses) and/or embryo (e.g. potyviruses) induce plantlet infection mechanically or systemically (Hamelin et al., 2017). More than

231 virus species are transmitted by seeds, mainly in Potyvirus, Nepovirus,

Cryptovirus, Ilarvirus, and Tobamovirus genera (Sastry 2013; Hull 2014). In addition to seeds, plant pollen has been confirmed as a source of transmission for several virus genera, including members of the genera Alphacryptovirus, Alfamovirus, Anulavirus,

Badnavirus, Cheravirus, Comovirus, Cucumovirus, Hordeivirus, Idaeovirus, Ilarvirus,

Nepovirus, Nucleorhabdovirus, Potyvirus, Sobemovirus, Tepovirus, Tobravirus,

Tymovirus, as well as viroids belonging to the genus of the Avsunviroid, Hostuviroid and even Pospiviroid (Das et al.,1961; George and Davidson, 1963; Gilmer and Way,

1963; Wang et al.,1993; Liu et al., 2014). Pollen is easily dispersed over a long distance by wind, honey bees and other pollen carriers like bumble bees, hoverflies, nectar scarabs and even thrips and are able to transmit some viruses to seeds and mother plants as well (Hull, 2014; Levitzky et al., 2019). Furthermore, dodder species

(Cuscuta spp.) parasitize many plants genera and are capable of transmitting plant viruses between adjacent plants as a bridge. Dodders are used as an experimental tool to transmit the viruses when they fail to be transmitted by other methods (largely mechanical) in the lab (Walkey, 1991). Some viruses may infect dodder cells and a low percent of seeds (Mikona and Jelkmann, 2010; Hull 2014).

Figure 1.5. The percentage of viruses that can be transmitted by different vectors (Syller 2014)

Based on the virus characteristics, being non-living obligate parasites, they often need to be acquired, transmitted and re-entered from infected plants to a living cell of new plants during an active process through vectoring by a diverse group of living organisms. Some species in arthropods (insects and mites), nematodes, fungi, and fungi-like organisms (protists zoosporic endoparasites) are among important so-called plant virus vectors. Mites, and mainly insects (aphids, thrips, whiteflies, leafhoppers, planthoppers, treehoppers, mealybugs and mirids) (Fig. 1.5) are the most common plant virus vectors. Nearly 450 aphid species that have a complex life cycle feed on sap of a diverse group of plants (about 5000 plant species), among them crop plants.

They are considered as crop pests to be the most important plant virus vectors as around 190 aphid species transmit over 300 virus species (of which 100 economically important ones) in different modes (persistent, semi-persistent and non-persistent)

(Emden and Harrington, 2017; Lacomme et al., 2017).

Figure 1.6. Stylet structure of aphids, Colored in yellow are maxillae, red are mandibels and green is labium. Adapted from (Taylor and Robertson, 1974; Uzest et al., 2010)

The complex structure of aphid mouth parts are specialized for sucking of plant saps

(Fig. 1.6). The two mandibles combine and form an outer stylet and maxillae join and piece to make inner stylet with canals including food and saliva ducts. These two canals join together and form common duct in the tip of the inner stylet that include a dense cuticular structure named acrostylet. This complex is wrapped in the labium at rest and when the aphid start feeding the sheath draw back and the stylet structure pierce the plant tissue as a needle (Harris and Maramorosch 1977). The saliva is injected from the saliva canal and plant sap is sucked up through the food canal.

Aphids inject two types of saliva, gel and watery, during plant sap sucking (Harmel et al., 2008). These two types of saliva involve in the different modes of virus transmission, as reviewed by Fereres (2007).

Aphids produce different morphological types during their life cycle (Moran, 1992;

Blackman and Eastop, 2000; Williams and Dixon, 2007). Most aphids may reproduce

both sexually and by parthenogenesis during their life cycle, which is called holocyclic, whereas some aphids are by parthenogenesis only. The aphids may produce various morphs throughout their life cycles including egg, nymphs, alate (winged or dispersing), apterous (wingless or non-dispersing) and brachypterous (having very short or incompletely-developed wings). Different environmental conditions trigger to change wingless to winged generation and vice versa. Among these triggers are aphid density, host plant quality (at least in some species), interspecific interactions, and environmental conditions (such as temperature, light and photoperiod) (cited in

Braendle et al., 2006). Some aphids (not all) produce winged forms (male and female) to move to new plants in case of crowding, nutritional and/or environmental pressures

(temperature, precipitation and light intensity) (Giordanengo et al., 2013).

Virus transmission by vectors as a specific molecular interaction show a range of species specificity. Some viruses can be transmitted by many vector species while some are only transmitted by a single vector species (Ng and Perry, 2004; Hogenhout et al., 2008; Bragard et al., 2013; Hull 2014; Syller, 2014).

Plant virus transmissions by insect vectors take place in four steps, including acquisition, latent period, retention, and inoculation. Specific variability in virus vector interaction result in a different transmission mode (Table 1.1). One of the important characteristics to categorize the mode of transmission is the virus retention site (stylet, foregut, gut and hindgut) and the route of virus circulation in the vector body (Watson and Robert 1939; Kennedy et al., 1962; Nault and Ammar 1989; Katis et al., 2007).

The most widely accepted criteria for classifying and categorizing virus transmission modes are based on the following factors:

 The Acquisition Access Period (AAP) or Acquisition Feeding Period (AFP) is

the time required between virus acquisition (by accessing or starting to feed)

and the ability of the virus transmission to new plants by vectors.The time

required between the access of the virulifer vector to plants and the inoculation

of the virus to plants is referred to as the Inoculation Access Period (IAP).

 The longevity of the retention of the virus (being a vector virulifer) ranges from

minutes to a lifetime, and can even include inheritance by the offspring.

 The location of retention and routing of the virus in the vector may be external

(surface cuticle) or internal (hemolymph and salivary gland) interactions.

 The molecular component of virus retention and replication in the body of

vectors.

 The latent phase of vectors is the interval between the acquisition of the virus

by vector from the infected plant and the ability to transmit the virus.

On the basis of the aforementioned parameters, the transmission mode of the virus is categorized by Hull (2014) as below:

 Non-circulative, non-persistent (NCNP)

 Non-circulative, semi-persistent (NCSP)

 Circulative, persistent; non-propagative (CNP)

 Circulative, propagative (CPP).

Table 1.1. Plant virus vectors and types of transmission.

Transmitted virus Mode of Vector References Family: Genera transmission Virgaviridae: (Pomovirus, Furovirus Protists Plasmodiophoral (rod-shaped), Pecluvirus) Campbell 1996; es: Benyviridae: Benyvirus zoospores Rochon et al., Spongospora, internally 2004; Bragard et Polymyxa Potyviridae: Bymovirus (filamentous) al., 2013

Varicosavirus (non-enveloped rod Campbell 1996; zoospores shape) Rochon et al., Aspiviridae: Ophiovirus 2004; Alfaro-

Fungi Olpidiaceae: zoospores Fernandez et al., Olpidium Tombusviridae: Carmovirus 2010; Bragard et Alphaflexiviridae: Potexvirus zoospores al., 2013; Hull (Filamentous) few species 2014 Rhizoctonia Bromoviridae: Cucumber mosaic Andika et al., Mycelium solani virus 2017 Longidoridae: van Hoof 1970, Longidorus, Karanastasi et al.,

Nematodes Secoviridae: Nepovirus, Cheravirus Paralongidorus 2000; Vellios et Xiphinema al., 2002; persistent MacFarlane, Trichodoridae: 2003; Andret-Link

Trichodorus, Virgaviridae: Tobravirus et al., 2004; Paratrichodorus Holeva and MacFarlane, 2006 Potyviridae: Tritimovirus, Rymovirus, Poacevirus Oldfield 1970; semi- Mites Kitajima et al., Eryophidae Betaflexivirida: Trichovirus persistent 2010, 2014; Alphaflexiviridae: Allexivirus Bragard et al., Secoviridae: Nepovirus (subgroup C) 2013;Solo et al., Rhabdoviridae: Dichorhavirus persistent- Teneopalpidae 2020; Fimoviridae: Emaravirus propagative Secoviridae: Comovirus, Gergerich, 2001;

(Beetles) Chewing Beetles: non-

insect: insect: Raccah and Chrysomellidae, Solemoviridae: Sobemovirus, circulative, Fereres, 2009; Coccinellidae, Tymoviridae: Tymovirus, foregut-borne Mukhopadhyay, Curculionidae Tombusviridae: Carmovirus (semi- 2011; Tolin et al., and Meloidae persistent) Bromoviridae: Bromovirus 2016; Pseudococcidae Closteroviridae: Ampelovirus (Mealybugs) and semi- Hull, 2014 Coccidaev (Soft Caulimoviridae: Badnavirus persistent scales) Caulimoviridae: waikaviruses and semi- badnaviruses persistent Sucking insect Sucking Geminiviridae: mastreviruses, and persistent Leafhopper curtoviruses circulative (Cicadelidae) Rhabdoviridae: Marafiviruses, Nucleorhabdovirus, Cytorhabdovirus, persistent- Ammar and Nault Phytoreovirus propagative 2002; Hull 2014;

Phenuiviridae: Tenuivirus Whitfield et al., Tymoviridae: Marafivirus 2015, 2018 planthopper Rhabdoviridae: Nucleorhabdovirus, persistent- (Delphacidae) Reoviridae: Fijivirus propagative Phenuiviridae: Tenuivirus planthopper persistent Nanoviridae: Nanovirus (Cixiidae) circulative

Ammar and Nault Treehopper Gemeniviridae:Topocuvirus, persistent 2002; Hull 2014; (Membracidae) Curtovirus, circulative Whitfield et al., 2018 semi- Secoviridae:Torradovirus, Jones 2005; persistent Czosnek and persistent- Gemeniviridae: Begomovirus, Ghanim 2012 circulative Whitefly Betaflexiviridae: Carlavirus, semi- Jeyanandarajah (Aleyrodoidea) Closteroviridae: Crinivirus persistent and Brunt 1993; Mansour and Potyviridae: Ipomovirus non-persistent Almusa 1993; Webb et al., 2012 propagative Rhabdoviridae: Nucleorhabdovirus Proeseler 1980 persistently Bugs: Odedara et al., Piesmatidae, Potyviridae: potyvirus non-persistent 2007 Miridae, Gibb and Randles Orsillidae (Lyga Solemoviridae: Sobemovirus persistent- 1988; Gibb and eidae: Orsillinae ( velvet tobacco mottle virus) circulative Randles 1991 ) and Potyviridae: Longan witches broom- circulative Pentatomidae Chen et al., 2001; associated virus Un-assigned mode in Seo et al., 2017 species salivary glands persistent- propagative, Bunyaviridae:Tospovirus transovarial Thrips transmitted (Thysanoptera) Jones, 2005 semi- Thripidae Tombusviridae: Machlomovirus persistent Solemoviridae: Sobemovirus pollen-carried Ilarviruses: Ilarvirus by thrips Betaflexiviridae: Carlavirus non-persistent non-circulative Bromoviridae: Alfamovirus, capsid Cucumovirus strategy semi- Caulimoviridae: Caulimovirus persistent helper strategy semi- Closteroviridae: Closterovirus persistent Comoviridae: Fabavirus non-persistent Ng and Perry Aphids 2004; Sanfacon et Luteoviridae: Enamovirus, circulative non (Aphididae) al., 2012; Bragard Luteovirus, Polerovirus propagative et al., 2013; Valli circulative et al., 2017 Nanoviridae: Nanovirus, Babuvirus non- propagative non-circulative Potyviridae: Macluravirus, Potyvirus helper strategy Rhabdoviridae: Cytorhabdovirus, circulative Nucleorhabdovirus propagative non-circulative Secoviridae: Waikavirus, Sequivirus semi- persistent

Non-circulative non-persistent transmission (NCNP)

NCNP viruses tend to infect all plant cell types during feeding and can be acquired from the infected plant and transferred to the recipient plant after AAP and IAP, respectively, within seconds to minutes by their vectors. These viruses are transmitted by the vector stylet when piercing the epidermal cells in order to find the desired plants and the vectors remain viruliferous for minutes to hours, depending on temperature and further feeding (Pirone, 1977; Roossinck, 2010). They lose their transmissibility as the vector molts or feeds on healthy plants for a few times (Nault and Styer, 1972, Powell and Hardie, 2000). During the virus transmission the virus particles don’t circulate (transit) across the vector body and the involved viral particles in the transmission are only retained in the tip of the insect stylet, and in the common duct (mainly aphid vectors). This may be mediated by interaction of the coat protein and vector receptor (Capsid-dependent) or by complex interactions, including coat protein, helper protein/s and vector receptor (Helper component mediated). Studies based on electrical penetration graph (EPG) and potential drop

(pd) waveform revealed that NCNP potyviruses are inoculated by aphid vectors during superficial brief intracellular punctures in the early host plant evaluation.

NCNP viruses acquire during sub-phases II-3 (third intracellular activity) and inoculate efficiently in sub-phases II-1 (first intracellular activity) during active saliva injection into the plant cells by aphids vectors (Martin et al., 1997; Powell, 2005;

Moreno et al., 2012). Alfamovirus, Bromovirus, Carlavirus, Cucumovirus, Fabavirus,

Macluravirus, Potexvirus, and Potyvirus are the most important NP viruses (Pirone and Perry, 2002; Bragard et al., 2013).

Persistent circulative, non-propagative transmission (CNP)

The persistently transmitted viruses, including Luteoviridae, Geminiviridae,

Nanoviridae, and Reoviridae, are transmitted by aphids, leaf- and treehoppers, and whiteflies. This type of transmission is more specific compared to non-persistently transmitted viruses. They are acquired and inoculated from and to the plant phloem within hours to days. Acquisition period can be roughly 5 min to hours, but at least a

12 h latency period is required from acquisition to inoculation, after which they can be inoculated in a timeframe of about 10-30 min, e.g. for aphid vectors transmitting luteoviruses (Hull, 2014). The retention time may last from a few days to the whole life span of the vectors, and may be retained even after molting. Virus particles must circulate and pass from the gut lining to the hemolymph and neighboring organs, and then to the accessory salivary glands of the vectors, after which the virus can be inoculated to the new plant host (Zhou, 2018). More than for the interactions between viruses themselves, plant and vector proteins, and sometimes extra proteins from other viruses (helper viruses) or even bacteria (Buchnera spp.) are needed to facilitate the transmission of these viruses (Gonçalves et al., 2005; Hull, 2014; Cilia et al., 2011;

2014), although the results of some studies raise doubts about the involvement of

Buchnera proteins in the transmission of Luteovirus (Bouvaine et al., 2011).

Non-circulative semi-persistent transmission (NCSP)

NCSP viruses possess features of the NCNP and CPNP viruses, and the virus particles circulating in the vector body and/or retained on the surface of the chitin lining at the tips of the stylet (common duct) or in the foregut of the insect vectors, are transmitted. The vectors do not transmit the virus after molting, and circulation of the virus through the body of the vector is not a requirement for transmission (Zhou, 2018).

They are often present in phloem, and AAPs, IAPs and retention periods (h to days) are typically longer than those of NP viruses. Details of this virus transmission mode are not well understood and there are still some obscurities (Childress et al., 1989;

Uzest et al., 2010; Chen et al., 2011; Ng, 2013; Li et al., 2016; Zhou et al., 2018).

Although some species of the genera Closterovirus, Crinivirus and Caulimovirus are

SP-type transmitted viruses, they use different sets of proteins and/or binding sites for transmission. Inoculation of NCSP viruses takes place during sub-phase II-2 in aphids, but they may also be transmitted by different vectors such as leafhoppers, aphids and whiteflies (Uzest et al., 2010; Moreno et al., 2012; Hull, 2014).

Persistent circulative, propagative transmission (CPP)

Propagative transmissible plant viruses enter and replicate in the insect vectors, and so, in this case, both plants and vectors may be considered to be hosts of the virus.

They are retained for days to a lifetime (even after molting vectors) and, depending on the virus, they infect various vector organs and even transmit transovarially to the offspring. This mode of transmission is extremely specialized and viruses are transmitted by a single species of insect vector. The propagative transmissible plant viruses are listed below. Marafivirus, Nucleorhabdovirus, Cytorhabdovirus, Tenuivirus,

Phytoreovirus, Fijivirus, and Oryzavirus are transmited by vectors belonging to the

Cicadellidae, Delphacidae, and Membracidae families, namely plant-, leaf- and treehoppers (Hohn 2007; Blanc et al., 2014; Hull ,2014; Whitfield et al., 2015; Dader et al., 2017).

1.2.2. Plant virus symptoms

Several hundreds of plant viruses on a vast number of plants with visible symptoms or invisible signs have been reported. Reports on virus presence without producing symptoms, so-called latent viruses, have been increasing since new techniques such as HTS became more commonly available in plant virus diagnostics (Adams et al.,

2018). The type and severity of symptoms are determined by type of virus (species and strains) and mainly the host plant (species, variety, resistance to the virus, virus

infection and plant age) (Kaplan and Meier, 1958; Funke et al., 2017; Ali and Abbo,

2019; Dupuis et al., 2019). Moreover, environmental conditions (Roossinck, 2015) and virus co-infection with other viruses or plant pathogens (Syller, 2012) affect the plant symptoms induction. Plant symptoms induced by viruses are yellowing and reddening, mosaic, stunting and dwarfing, distortion, crinkle, flower color breaking, local lesions and ring spots, vein clearing and necrosis, vein banding, hypertrophy and hypotrophy, leafroll and leaf curl, epinasty and big-vein (Agrios, 2005; Hull, 2014).

1.2.3. Important potato viruses

Vegetative propagation of potato and accumulation of different viruses in tubers degenerate seed tubers and impose high costs to virus elimination by means of some methods like tissue culture (Thomas-Sharma et al., 2015). About 50 virus species infect potato and cause an important crop losses, however, they rarely kill the potato plants (Wale et al., 2008). The reported naturally occurring plant viruses (and viroids) in potato worldwide are listed in Table 1.2.

Table 1.2. The viruses infecting potato (S. tuberosum)

Name Taxonomy Losses No. Distribution Vector References

(Abbreviation) (Order, Family, Genus) significance

Unassigned, Little Alfalfa mosaic Stevenson et al., 1 Bromoviridae, Worldwide economic Aphid virus (AMV) 2009 Alfamovirus importance

Andean potato Tymovirales, Koenig et 2 latent virus S America Little damage Beetles Tymoviridae, Tymovirus al.,1979 (APLV)

Andean potato Picornavirales, may be Dusi and Avila, 3 mottle virus Secoviridae, S America Contact significant 1988 (APMoV) Comovirus

Arracacha virus Cheravirus, (tentative), Jones, 1981 4 Peru, Bolivia Unknown Unknown B-oca strain Sequiviridae

(AVB-O)

Unassigned, Little Beet curly top Arid areas 5 Geminiviridae, economic Leafhoppers King et al., 2012 (BCTV) Worldwide Curtovirus importance

Cucumber Unassigned, Little Chrzanowska et 6 mosaic virus Bromoviridae, Worldwide economic Aphid al., 2003 (CMV) Cucumovirus importance

Eggplant mottled Mononegavirales, Rare in 7 dwarf virus Rhabdoviridae, Iran Aphid Danesh, 1989 potatoes (EMDV) Nucleorhabdovirus

Groundnut bud Unassigned,

8 necrosis virus Bunyaviridae, India Unknown Thrips Jain ey al., 2004

(GBNV) Tospovirus

Groundnut Unassigned, Bragard et al., 9 ringspot virus Bunyaviridae, Argentina Unknown Thrips 2020 (GRSV) Tospovirus

Impatiens Unassigned, Crosslin and

10 necrotic spot Bunyaviridae, USA Unknown Thrips Hamlin, 2010

virus (INSV) Tospovirus

Potato aucuba Tymovirales, Contact, Susaimuthu et 11 mosaic virus Alphaflexiviridae, Worldwide Unknown aphids al., 2007 (PAMV) Potexvirus Picornavirales, Potato black Richards et al., 12 Secoviridae, S America Unknown Unknown ringspot virus 2014 Nepovirus Potato black Picornavirales, Low Salazar and 13 ringspot virus Secoviridae, Peru importance in Nematodes Harrison 1978 (PBRSV) Nepovirus potato Potato deforming Geminivirus, Bragard et al., 14 mosaic Brasil Up to 35 % Whitefly Begomovirus, tentative, 2020 (Argentina) (PDMV) Potato latent Tymovirales,Betaflexiviridae, 15 N.America Unknown Aphid Nie, 2009 virus (PotLV) Carlavirus Potato leaf Unassigned, De Boer et 16 Worldwide up to 90% Aphid rollvirus (PLRV) Luteoviridae, Polerovirus al.,1996 Unassigned, Potato mop top N and C Affect tuber Stevenson et al., 17 Virgaviridae, Fungi virus (PMTV) Europe Peru quality 2009 Pomovirus Potato rough Argentina Little Massa et al., 18 dwarf virus Tentative, Carlavirus Aphid Uruguay importance 2008 (PRDV)

Contact and Potato spindle Unassigned, 64 % Pfannenstiel, aphid when 19 tuber viroid Pospiviroidae, Europe (Pfannenstiel, 1980 Coinfected (PSTVd) Pospiviroid 1980) with PLRV Unassigned, Potato virus A Can be up to De Boer et 20 Potyviridae, Worldwide Aphid (PVA) 40 % al.,1996 Potyvirus Tymovirales, Potato virus M De Boer et 21 Betaflexiviridae, Worldwide at worst 15 -45% Aphid (PVM) al.,1996 Carlavirus Tymovirales, 20 - 80 % Potato virus P Bragard et al., 22 Betaflexiviridae, Brasil importance Aphid (PVP) 2020 Carlavirus local Tymovirales, Potato virus S At worst 10 - De Boer et 23 Betaflexiviridae, Worldwide Aphid (PVS) 20 % al.,1996 Carlavirus Tymovirales, Potato virus T S America Bragard et al., 24 Betaflexiviridae, Unknown Contact (PVT) (Peru) 2020 Unassigned Picornavirales, Potato virus U Bragard et al., 25 Secoviridae, Peru Unknown Nematodes (PVU) 2020 Nepovirus Unassigned, Potato virus V N.America Damage Oruetxebarria et 26 Potyviridae, Aphids (PVV) S.America severe at. 2000 Potyvirus Tymovirales, Potato virus X Usually De Boer et 27 Alphaflexiviridae, Worldwide Contact (PVX) 15 -20% al.,1996 Potexvirus Unassigned, Potato virus Y Losses reach Aphid Bragard et al., 28 Potyviridae, Worldwide (PVY) 10-80 % 2020 Potyvirus Potato yellow Mononegavirales, No economic Anderson et al., 29 dwarf virus Rhabdoviridae, N.America Leaf hopper importance 2018 (PYDV) Alphanucleorhabdovirus Potato yellow Unassigned, Carribean Morales et al., 30 mosaic virus Geminiviridae, Unknown Whitefly region 2001 (PYMV) Begomovirus S America Potato yellow Unassigned, (Columbia Salazar et al., 31 vein virus Closteroviridae, More than 50 % Whitefly Venuzuela 2005 (PYVV) Crinivirus Peru and Potato yellowing Silvestre et al., 32 Tentative, Alfamovirus S America Unknown Aphid virus (PYV) 2011 Bragard et al., Potato yellow Unassigned, 2020; Nisbet et 33 blotch virus” Potyviridae, UK Unknown Aphid al., 2018; (PYBV) Potyvirus Kreuze et al., 2020 Solanum apical Significance in Hooker and 34 Tentative, Geminivirus Peru Aphid leaf curling virus localized Salazar 1983

(SALCV) areas

Sobelivirales, Bragard et al., Sowbane mosaic Rare in 35 Unassigned, Worldwide Unknown 2020; Kreuze et virus (SoMV) potatoes Sobemovirus al., 2020 Tobacco Unassigned, Bragard et al., Argentina 36 chlorotic spot Bunyaviridae, Unknown Unknown 2020; Kreuze et Brazil virus (TCSV) Tospovirus al., 2020 Unassigned, Tobacco mosaic Not problem in 37 Virgaviridae, Worldwide Contact Jung et al., 2002 virus (TMV) potatoes Tobamovirus Tobacco Unassigned, Europe, Not Olpidium Beuch et al., 38 necrosis virus Tombusviridae, N.America, significance in brassicae 2013 (TNV) Necrovirus Tunisia potatoes Unassigned, Tobacco rattle May be David et al., 39 Virgaviridae, Worldwide Nematodes virus (TRV) appreciable loss 2010 Tobravirus Unassigned, Little Tobacco streak Salazar et 40 Bromoviridae, S America significance in thrips virus (TSV) al.,1982 Ilarvirus potatoes Picornavirales, Tomato black Little 41 Secoviridae, Europe Nematodes Kaiser, 1980 ring virus (TBRV) significance Nepovirus Tomato leaf curl Unassigned, Usharani et al., 42 New Delhi virus Geminiviridae, India Unknown whitefly 2004 (ToLCNDV) Begomovirus Unassigned, Yazdani- Tomato mosaic Not problem in 43 Virgaviridae, Hungary Contact Khameneh et virus (ToMV) potatoes Tobamovirus al., 2013 Tomato mottle Unassigned, Cordero et al., 44 Taino virus Geminiviridae, Cuba Unknown Whitefly 2003 (ToMoTV) Begomovirus Unassigned, Tomato spotted Worldwideh Importance in Abad et al., 45 Bunyaviridae, Thrips wilt virus (TSWV) ot climates Localized areas 2005 Tospovirus Tomato yellow Bunyaviridae, Golnaraghi et 46 fruit ring virus Iran Unknown Thrips Tospovirus al., 2008 (TYFRV) Tomato yellow Bunyaviridae, Birithia et al., 47 Poland Unknown Thrips ring virus (TYRV) Tospovirus 2012 Tomato yellow Unassigned, Ribeiro et al., 48 vein streak Geminiviridae, Brazil Unknown Whitefly 2006 virus(ToYVSV) Begomovirus Wild potato Unassigned, Potyviridae, No problem in Fribourg et al., 49 mosaic virus - Aphids Potyvirus potatoes 2019; (WPMV) Pepino mosaic Tymovirales, Not infected Bragard et al., 50 virus Alphaflexiviridae, - Contact potato naturally 2020 (PepMV) Potexvirus

Even though about 50 distinct viruses and one viroid have been recorded worldwide to infect potatoes (Table 1.2.), internationally, only a few of them inflict substantial losses, including: Potato leafroll virus (PLRV), Potato Virus Y (PVY), Potato Virus A

(PVA), Potato Virus S (PVS), Potato Virus M (PVM), Potato Virus X (PVX), Potato aucuba mosaic virus, Potato mop-top virus, Alfalfa mosaic virus (AMV). PLRV is the most important potato virus in North America whereas PVY is the most prevalent and important virus in the EU. These two viruses may reduce potato yield up to 75% whereas others can reach 30% losses (Stevenson et al., 2009; Rudelsheim and

Smets, 2012). Many potato viruses are transmitted by aphids in persistent and non- or semi-persistent manners, and as such they are the most important virus vectors in potato. The most important and common in potato viruses worldwide are discussed individually in the following paragraphs.

Potato virus Y (PVY)

PVY is the type species of Potyvirus genus in Potyviridae family and is one of the most important potato viruses. PVY is considered as the 5th most important plant virus worldwide. It has a relatively wide host range and a significant economic impact on many commercial crops (mainly solanaceous), explaining the PVY importance

(Scholthof et al., 2011). PVY was first described by Smith (1931) and its importance has increased along with increasing potato cultivation. PVY infections decrease the quality and quantity of potato yields and result in seed degeneration, which imposes huge economic losses and extra costs in potato cropping and seeds production. PVY infection on potatoes causes a variety of symptoms (from mild to severe), including yellows, mottling, mosaic, rogues, yellows, necrosis, leaf malformation, plant defoliation, and potato tuber necrotic ringspot diseases (PTNRD) (Fig. 1.7). The intensity and type of these symptoms also vary depending on environmental

conditions and genetic diversities of both PVY (the virus strain) and the host plant (e.g. potato cultivar) (Kerlan 2006; Nie et al., 2012; Lacomme et al., 2017). Economic impact of PVY on potato in different countries are reported as 16.5% in Ireland, 34% in Canada, 37% in Kenya, 40-44% in Poland and the USA and about 50% in China

Figure 1.7. Symptoms of PVY; a range of Mosaic, Tissue necrosis and PTNRD Cornell University. (https://blogs.cornell.edu/potatovirus/pvy/pvy-symptoms-and- diagnosis)

(Gray et al., 2010; Wang et al., 2011; Were et al., 2013; Hasiow-Jaroszewska et al.,

2014; Hutton et al., 2015).

Potato leafroll virus (PLRV)

PLRV is a phloem limited +ssRNA virus member of the genus Polerovirus with virions of icosahedral structure. PLRV is transmitted and established in the field by growing infected tubers and is transmitted in CPNP by aphids (mainly M. persicae) with a latent period of 1–2 days. PLRV and PVY are the most important potato virus diseases and have been reported in potato fields worldwide. Infected plants, depending on host cultivar and virus strain, show leaf rolling, yellows, purplish discoloration, plant dwarf and tuber necrosis (Fig. 1.8) (Wale et al., 2008).

Figure 1.8. Symptoms of PLRV (CABI, https://www.cabi.org/isc/datasheet/42783)

Potato virus S (PVS)

PVS is a Carlavirus (Flexiviridae) with a +ssRNA genome and filamentous structure.

The virus is transmitted mechanically and by several aphid species in NCNP manner including M. persicae, Aphis nasturtii, Rhopalosiphum padi and Aphis fabae (Santillan et al., 2018). Infected plants mostly show mild or no symptoms, yet this depends on the strain (PVSA and PVSO) and plant cultivar. Infected leaves show bronzing, are rough when touched, produce necrotic spots, often the tips are inclined downwards

(Fig. 1.9) and plants generally remain smaller. PVS is not a serious disease, except when it occurs in coinfection with other viruses (Loebenstein and Gaba, 2012).

Figure 1.9. PVS symptoms (INRA, http://ephytia.inra.fr/en/C/21030/Potato-Symptoms)

Potato vrus M (PVM)

PVM is a member of the genus Carlavirus and has a +ssRNA genome. It is known as a prevalent and economically important potato virus with crop losses ranging from 15 to 45%. Occasionally, even 100% of the plants in a plot may be infected. In potato plants, PVM causes mottling, mosaic, crinkling and rolling of the leaves, and plant dwarfing (Fig. 1.10). In addition to mechanical transmission, PVM is transmitted by aphids in a non-persistent manner (Wetter, 1972).

Figure 1.10. Rolling of the potato leaves infected by PVM (INRA, http://ephytia.inra.fr/en/C/21040/Potato-Potato-virus-M-PVM)

Potato virus X (PVX)

PVX is a mono-partite filamentous +ssRNA virus belonging to the Potexvirus genus, that causes a range of symptoms from severe leaf mosaic and distortion to completely

symptomless in potato (Fig. 1.11), resulting in yield losses of about 10 to 20%. The strains of PVX are readily sap-transmissible and are mainly transmitted through mechanical contact in nature. However, PVX is also transmitted by aphids when it is co-infected with other potyviruses PVY or PVA (Govier and Kassanis 1974a, 1974b;

Koenig and Lesemann, 1989).

Figure 1.11. Mosaic on potato leaves caused by PVX. (INRA, http://ephytia.inra.fr/en/C/21026/Potato-Symptoms)

Potato yellow blotch virus (PYBV)

PYBV is a recently described rare species of the genus Potyvirus infecting potatoes.

It produces isolated yellow blotches as most striking leaf symptoms, and based on sequence analysis and host range, it is closely related, yet distinct, from potato virus

Figure 1.12. The symptoms of PYBV on potato leaves

A (PVA) (Fig. 1.12) (Nisbet et al., 2018). PYBV possesses a +ssRNA genome and is likely to be spread by tubers and aphid vectors in an NCNP manner, like other virus species in the same family, however, transmission assays are needed to confirm this.

Wild potato mosaic virus (WPMV)

WPMV is a distinct species that holds all general characteristics of potyviruses

(+ssRNA and NCNP transmission by aphids) (Jones, 1979), and is closely related to

PVY and peru tomato virus (PTV), PVV, pepper mottle virus (PepMoV), pepper severe mosaic virus (PepYMV) and pepper yellow mosaic virus (PepYMV). These viruses are suggested to form an individual clade within the group of the potyviruses (Spetz et al.,

2003; Spetz and Valkonen, 2003; Fribourg et al., 2019). WPMV has been detected in and observed to induce mosaic in 16 wild tuber forming Solanum spp., in South

America. However, none of the 13 potato cultivars (S. tuberosum) that have been tested resulted in infection (Jones, 1979).

Potato virus V (PVV)

PVV was first reported as a mutant of PVYC by Rozendaal et al., (1971), and later in

1984 described as a distinct virus (Jones and Fuller 1984). Nowadays PVV accounted for 4% of the total potato viruses in UK (Pickup et al., 2009). Depending on the potato cultivar, it is showing mosaic, crinkling and systemic necrosis in naturally occurring infections. Mechanical inoculation induces local lesions and hypersensitivity (Fig

1.13). Like other potyviruses, PVV consists of a +ssRNA genome and is transmitted by aphids in an NCNP manner (Jones and Fribourg 1986; Shamsadden-Saeed et al.,

2013).

Figure 1.13. Mild mosaic vein clearing, crinkling, systemic necrosis

Potato virus A (PVA)

PVA is one of the most important potato viruses worldwide. Depending on the virus strain, potato cultivar and time of infection, infected PVA potatoes may show no visual symptom, or a range of symptoms on leaves including slight blurred border mosaic and mottling without distortion (Fig. 1.14). Different aphid species transmit PVA

(+ssRNA) in an NCNP manner. PVA causes crop loss in potatoes by up to 40%

Figure 1.14. PVA symptoms on potato (INRA, http://ephytia.inra.fr/en/C/21041/Potato-Symptoms)

(Kreuze et al., 2020). Co-infection of PVA with PVY and/or PVX may increase symptom severity to crinkling and severe leaf deformation, resulting in raised yield

losses (Bartels, 1971).

1.3. Potato virus Y (PVY) and its characteristics

1.3.1. Taxonomic classification

PVY is the type member of potyvirus genus in potyviridae family (Fig. 1.4). Potyviridae is the second largest plant virus family (after Geminiviridae) and includes 12 genera

(Arepavirus, Bevemovirus, Brambyvirus, Bymovirus, Celavirus, Ipomovirus,

Macluravirus, Poacevirus, Potyvirus, Roymovirus, Rymovirus and Tritimovirus).

Potyviruses underwent a radiative evolution in 7,000 years, at the same time as agricultural intensification in the middle of Holocene (Moury and Desbiez 2020). The virion shape is filamentous and non-enveloped consisting of one (lengths: 650-900 nm), or two (lengths: 500-600 and 200-300 nm) particles (12-15 nm in diameter). The genome of the viruses in this family are linear +ssRNA monopatite, except for bymoviruses that consist of bipartite genomes with two ORF (Fig. 1.15).

The Potyvirus genus as the second largest plant virus genus (after Begomovirus with

424 species), with 183 species and is the most largest, distributed and economically important RNA virus genera worldwide in cropping ecosystems (Genus: Begomovirus,

2021; Genus: Potyvirus, 2021). Despite the large number of potyviruses, only five

Figure 1.15. Genome organization (mono and bi partite) in the family Potyviridae. (Adapted from: https://viralzone.expasy.org/48?outline=all_by_species)

species are known to naturally infect potatoes; potato virus V (PVV), potato virus A

(PVA), potato yellow blotch virus (PYBV), wild potato mosaic virus (WPMV) and potato virus Y (PVY) (Table 1.2). They show a high similarity in host range, serology, and sequence data compared to other potyviruses (Spetz et al., 2003; Spetz and

Valkonen, 2003).

1.3.2. Genome organization and proteins

As other potyviruses, PVY is filamentous, and possesses flexuous virus particles (700 nm x 11-13 nm) and a positive sense single stranded RNA genome (+ssRNA), encapsidated in a coat protein with 2000 unites of same monomer (30 kDa). The RNA genome length is about 9700 nucleotides ending with a viral protein genome-linked

(VPg) at the 5’end and a poly-A tail in 3' end (Fig. 1.16).

Moreover, VPg is a multifunctional protein at the 5′ terminus of +RNA and acts mainly as a primer during RNA synthesis in a variety of some +ssRNA viruses, including

Potyviridae wherase poly-A in 3' of RNA terminus (mRNA and +RNA viruses) that has only adenine bases and is involved in gene expression. The PVY genome contains two open reading frames (ORF), including a main ORF that is translated to a large polyprotein that breaks down into 10 functional proteins, and the small ORF named pretty interesting Potyvirus ORF (P3N-PIPO or simply PIPO) that is a +2 frame shift within the main ORF and is translated to a small protein (Table 1.3) (Hu et al., 2011;

Cuevas et al., 2012; Hillung et al., 2013; Revers and García, 2015; Lacomme et al.,

2017; Valli et al., 2018).

Figure 1.16. Genome Diagram of PVY (describe in table 3) (Cuevas et al., 2012)

Table 1.3. Genome and functions of Potyvirus proteins (Hu et al., 2011; Hillung et al., 2013; Revers and García, 2015; Lacomme et al., 2017; Valli et al., 2018)

Gene nucleotide, Protein size Proteins Function size (bp) (aa) 5’ UTR 5′ untranslated 1-188, 188 - Regulation of translation region Serine proteinase, Accessory factor for virus amplification, Host adaptation, Stabilizes CP, P1 189-1013, 825 275 stimulate HC-pro silencing suppression, Binds RNA. Cysteine proteinase, Helper factor for aphid transmission, RNA silencing suppression, HC-Pro 1014-2408, 1395 465 Enhancement of yield of virus particles, HC proteinase, Resistance breaking, Virus movement. Virus amplification, Host adaptation and P3 2409-3503, 1095 365 pathogenicity, Resistance breaking

P3N-PIPO 62–76 Viral cell to cell movement.

Modulation of P3 activity? Exact function is 6K1 3504-3659, 156 52 unknown. Role in potyviral infection. RNA helicase, RNA replication, Pinwheel formation (Forms the cylindrical cytoplasmic CI 3660-5561, 1902 634 inclusions), Virus movement, Cell to cell movement, Binds RNA. Membrane vesicles proliferation, Membrane 6K2 5562-5717, 156 52 targeting to ER-types membrane, Replication, Systemic movement. Genome-linked protein, Primer of RNA replication, RNA translation, Cell-to-cell and systemic Virus VPg 5718-6281, 564 188 movement, Nuclear Inclusion, Nuclear transport, Resistance breaking, Symptom modification. Cysteine proteinase, DNAse, Small Nuclear Inclusion protein A, Serine-like proteinase activity: Cleave P3-6K1-CI-6K2-Nia-Nib-CP, PrimeRNA NIa 6282-7013, 732 244 synthesis, Required for systemic infection, Nuclear transport, Cell-to-cell and systemic movement, Resistance breaking. Large Nuclear Inclusion protein B, RNA replicase, NIb 7014-8570, 1557 519 Viral replication, Nuclear transport, Symptom

modification. Coat Protein (Protection of genomic RNA), Cell-to- cell and systemic movement, Aphid transmission, CP 8571-9371, 801 267 Virus encapsidation and Virus assembly, Regulation of viral RNA amplification, Seed transmission, Symptom modification, Translation. 3’ UTR 9372-9702, 331 - - Full length 1-9702, 9702 3061 -

1.3.3. Genetic diversity and strains

Genetic variation in RNA viruses (as well in PVY) known as quasispecies and strain demarcations originate from mutations, recombination (and reassortment in multipartite viruses) and selection pressures from host, virus and presumably vector interaction (Schneider and Roossinck, 2001; Tromas and Elena, 2010 Nanayakkara et al., 2012a; Kutnjak et al., 2015; Domingo and Perales, 2019). Like other +ssRNA viruses, PVY genome is a functional mRNA (as a basic strategy of virus replication) and is translated shortly after penetration into the host cell. This strategy (fast translation and replication) together with lack of proofreading activity of RNA virus polymerases (Elena and Sanjuán, 2005; Elena et al., 2011), might be the reasons for the high mutation rate and genetic diversity in the +RNA viruses in general, and PVY in particular (Wolf et al., 2018). As an outcome of high mutation rate, the RNA viruses are more adaptive to changes of conditions in comparison with DNA organisms, including DNA viruses (Duffy, 2018). However, random genetic drift during horizontal transmission of these RNA viruses by sucking vectors like aphids, may decrease diversity and could be important in virus evolution (Betancourt et al., 2008).

PVY as a complex of different strains, based on TMRCA (time to most recent common ancesto) and using the Bayesian tip-dating of the non-recombinant dated world PVY population, originated from southern America around 156 CE (1841 and 1879 years

before 2016), where also the potato originated. Along with potato, as a new plant, PVY was introduced to Europe in the 16th century by infected tubers. The most genetic diversity followed by breeding programs to manage late blight epidemics in mid-19th century in Europe (Fuentes et al., 2019; Green et al., 2020). Breeding programs and growing PVY resistance cultivars, intensive potato cropping and pesticide application changed natural prevalence of resistance genes pool and the PVY transmission types

(by aphid, tuber or mechanical) compared to natural agroecosystems. These factors and their results in turn affected the natural selection of emerging mutants in the new conditions and the PVY strain diversity (Funke et al., 2017; Dupuis et al., 2019; da

Silva et al., 2020). PVY strains were first classified, based on the type of PVY symptoms in tobacco cultivars, the response of potato cultivars carrying N genes (Ny,

Nc, Nz, Ny-1and Ny-2) for necrosis and hypersensitive response (HR) and the induction of PTNRD in potato cultivars (Szajko et al., 2008, 2014). Different PVY proteins induce plant N genes to initiate HR, e.g. HC and NIa proteins that elicit Ny and N genes to induces HR, respectively (Singh et al., 2008; Chikh Ali et al., 2013;

Chikh Ali et al., 2014; Kehoe and Johnes, 2015; Baebler et al., 2020). Currently, PVYO,

PVYN, PVYE, PVYC, PVYZ, PVYNTN and PVYN-Wi are the main recognized biological strain groups (path groups). Kehoe and Jones (2015), however, also consider a PVYD strain group eliciting an Nd gene. The PVYO, PVYC and PVYZ strains trigger HR in potato cultivars carrying Nytbr, Nctbr and Nztbr genes, respectively, whereas PVYN and PVYE do not induce HR in the presence of these potato genes. Instead they cause vein necrosis and mosaic in tobacco respectively. All strain groups, except for PVYC, are able to induce PTNRD, depending on the potato cultivar and environmental conditions, in particular temperature (Le Romancer and Nedellec 1997). PTNRD was induced by different PVY strains on a range of susceptible cultivars after mechanical

inoculation experiments under controlled environmental conditions. However, PVYNTN is known to be responsible for the most of the reported PTNRD cases in most of the cultivars (Gibbs et al., 2017; Green et al., 2017). The symptoms and interaction of PVY strain groups and potato genotypes are summarized in Table 1.4. Despite being contaminated with PVYNTN isolate, Le Romancer and Nedellec (1997) found that certain cultivars, such as Spunta, Maris Piper, and Thalassa, did not develop PTNRD.

This finding demonstrated that at least one major dominant severe resistance gene

(Ry) regulates this reaction in non-sensitive cultivars, resulting in strong resistance to

PTNRD and tuber necrosis prevention in field conditions.

In addition, serological approaches have been used as classical techniques for the detection and discrimination of PVY serotypes. Two main serotypes are described using the immunological properties of PVY isolates. The main N serotype included

PVYN, PVYNTN, PVYZ and PVYE, and the main O/C serotype included PVYO, PVYC and PVYN-W. PVY serotyping illustrates the variation of the coat protein, that is a small part of the genome, and a point mutation may alter the serotype without changing the pathological features (Chikh Ali et al., 2007a). The diversity of mono- and polyclonal antibodies and also detection of different strains belonging to the same serotype

(N=NTN and NWi=O) restrict the usefulness of serological methods (Singh et al.,

2008). Biological and serological properties attribute to small differences in the nucleic acid sequences and, in some cases, these classification systems are not compatible

(Blanco-Urgoiti et al., 1998; Kehoe and Jones, 2015).

Since the 1980s, the key strategy in PVY strain differentiation was the available genome sequences and more in particular, the recombination patterns (Carrington and Dougherty, 1987; Robaglia et al., 1989). However, major biological variations or serological properties (host range or symptoms, and also strain group) were also still

considered. New recombinants have recently been detected, and more recombinants continue to emerge (Chikh Ali, 2007b; Chikh Ali, 2010a, b; Green et al., 2020).

Table 1.4. Symptoms of PVY strain groups on Nicotiana tabacum and potato cultivars with different genotypes. Potato Tuber Necrotic Ringspot Disease (PTNRD), (og) only in greenhouse, (HR) hypersensitive response, (VN) veinal necrosis, (Mo) mosaic, (S) susceptible, * hypothetical Suggested new PVY strain (PVYD) that elicit a new resistance gene (Ndtbr), - no information. (Jones, 1990; Kerlan and Tribodet, 1996; Valkonen, 1997; Kerlan et al.,1999, 2011; Browning et al., 2004; Piche et al., 2004; Szajko et al., 2008, 2014; Barker et al., 2009; Hu et al., 2009; Gray et al., 2010; Galvino-Costa et al., 2012; Karasev and Gray, 2013; Chikh Ali et al., 2014; Tomczynska et al., 2014; Glais et al., 2015; Kehoe and Jones, 2015; Rowley et al., 2015).

Plant Genotype of Solanum tuberosum species Frequency of Nicotiana PTNRD in Nztbr, Ny-1, PVY tabacum Nytbr Nctbr Ndtbr* potato Nctbr,Nytbr Ny-2 strains cultivars groups O Mo HR S HR HR - ++- (og) C Mo S HR HR - - - N VN S S S HR - ++- (og) Z Mo S S HR - - ++- E Mo S - S - - ++- N-Wi VN S S S HR - ++- (og) NTN VN S S S HR - +++ D* - - - - - HR ?

Genomic studies reveal a big variability in PVY populations (Kehoe and Jones, 2015;

Green et al., 2017; Fuentes et al., 2019), as Kehoe and Jones (2015) divided PVY strains to 13 phylogroups. Based on genome sequences and recombination patterns of about 400 isolates, 5 strains, namely PVYO, PVYO-O5 (or PVY-O5), PVYEU-N, PVYNA-

N and PVYC are considered as non-recombinant, and thirty six strains are the recombinants originating from these 5 non-recombinant strains (Green et al., 2017)

(Fig. 1.17). However, Green et al., (2020) classified isolates from across the world to

6 minor phylogroups PVYO-O5, PVYO-O (YZ-CM1), PVYEu-N, PVYEu-N (YN-RM) PVYNA-N

(YN-M) and PVYC1 (YC-R) and at least fourteen recombinant phylogroups.

PVYO-O5 and PVYNA-N are O and N strains, respectively, that are prevalent in North

America. Fuentes et al., (2019) believed that PVYN originated from potato in South

America, but the origin of the O and C strains are unknown. The recombinant strains have been evolving and distributed in Europe by new potato cultivars seeds that were bred to manage older PVY strains including non-recombinant and thence, new cultivars forced to select and increase population of the some mutate PVY strains (as natural pressure selection) (Funke et al., 2017; Dupuis, 2019; Karasev, 2019).

Diverse strains are dominant in various geographical regions, which can depend on the abundance and background of prevalent cultivars. Strains N-Wi and NTN 1 are most prevailing in the USA, respectively (Karasev et al., 2011; Funke et al., 2017).

The PVYNTN is also the most prevalent strain group in Switzerland, even though it has decreased from 78.9% in 2003 to 71.0% in 2012 (Dupuis et al., 2019 ). Studied PVY strains population in German potato farms from 2011 to 2013 revealed that PVYO abundance has dropped from 63% to 15% whereas the PVY recombinant has increased from 34% to 61% in which PVYN-Wi (60%) and PVYNTN (40%) became prevalent recombinants (Lindner et al., 2015).

Fig. 1.17. Types of PVY recombinants (Green et al., 2017).

Recent studies reveal PVYEu-NTN and PVYN:O are the most predominant strains in

Canada (MacKenzie et al., 2018) and NTN is the most important strain in Australia

(Rodriguez-Rodriguez et al., 2020). A study, conducted in the Middle East by Chikh-

Ali and colleagues (2008), as well as Hosseini et al. (2011), reveal that characterized new recombinant PVYNTN-NW-SYR-I and PVYNTN-NW-SYR-II are dominant strains in

Syria, while a closely related strain to the European PVYNTN is more common in Iran.

Traces of the Syrian PVY strains in Chinese potato farms is proven by Bai et al. (2019) recently. They showed that PVYNTN-NW-SYR-II and PVYN-Wi both are the most prevalent strains in China, followed by PVYNTN-NW-SYR-I, PVYN:O, and Eu-PVYNTN.

Serological tests were used by Rolot and Steyer (2008) and Rolot (2009) to determine the important PVY strains in Belgium. The N group was reported to be the most prevalent PVY strain group in potato farms. PVY was observed in 60-80 percent of the fields tracked from 2009 to 2011, according to a new report by Bosquée et al. (2016).

In 2010, the incidence rate in farms with seed and stock potatoes was 21.0 percent and 62.0 percent, respectively. They found that the N/NTN strain of PVY predominated in 70-100 percent of the checked samples, with the exception of potato seed farms in

2011, which had 72.73 percent of the O strain. Despite using Wilga strain specific primers, they were unable to detect it in Belgian potato farms (Bosquée et al., 2016).

1.3.4. Host range

A wide diversity of plants are listed as natural PVY hosts, and comprise of 495 species in 72 genera of 31 families. It includes important edible crops such as potato, tomato, pepper, tobacco and eggplant, in addition to 211 species within 9 genera of the

Solanaceae, putting this family on top of the host list of PVY. Moreover,

Amaranthaceae, Fabaceae, Chenopodiaceae, Compositae and Brassicaceae are families that include numerous other important host species (Kerlan, 2006; Kaliciak

and Syller, 2009). Some plants like Solanum nigrum, Hibiscus trionum, Amaranthus retroflexus and Physalis sp. are important weeds as reservoirs of PVY also being preferred hosts for PVY vectors in agricultural farms (Arli-Sokmen et al., 2005; Chikh

Ali et al., 2008; Cervantes and Alvarez, 2011). Further study showed that weeds such as Erodium cicutarium, Geranium pusillum, Lactuca serriola and Lamium purpureum could be infected by PVY but do not show visible symptoms (Kazinczi et al., 2004).

The huge host diversity could indicate different evolutionary routes, increasing the

PVY strain diversity by means of natural selection pressure, together with other diversity factors such as mutation and recombination (Garcia-Arenal and Fraile, 2013).

In addition, the incidence of virus diseases can be affected by the number and diversity of plant hosts of virus, and therefore, epidemiology and management strategies of the virus disease could be more complicated and lead to increasing costs, particularly when weeds play an important role in the crop system (Ormeno et al., 2006; Moury and Desbiez, 2020).

1.3.5. Transmission

PVY is transmitted from infected plants to offspring in the next generation (vertically) sexually (true seeds) or vegetatively (tubers) and from infected plants to the plants of the same generation (horizontally), mainly by aphids and mechanical contacts of adjacent plant organs (roots or foliage). PVY-infected tubers and aphid transmission are the main ways of PVY natural dispersal. In addition, plants grown out of PVY infected tubers lead to plants growing more slowly, resulting in a lower yield (da Silva et al., 2020; Hegde et al., 2020).

Different aphid species transmit PVY in NCNP manner at a different relative transmission efficiency (Lacomme et al., 2017). Transmission efficiency is a complex phenomena that are influenced by different factors. The most important factors that

have been discussed in PVY transmission are related to: 1) host plant source of the virus (source of virus acquiring); 2) transmissibility of virus strains or isolates; 3) the host plant that is receiving the virus from the aphid vector (inoculated plants); 4) the plants on which the aphids were feeding (reared) before virus acquisition (Al-Mrabeh,

2010; Bosquee et al., 2018); and 5) aphid species or clones. Moreover, PVY transmission is also affected by environmental conditions such as atmospheric gas concentration (Dader et al., 2016; Bosquee et al., 2018), temperature, and (relative) humidity (Singh et al., 1988; Qamar et al., 2003; Chung et al., 2016). These environmental factors could change virus transmission as outcome of changing aphid behavior, plant susceptibility and/or virus replication in plants (Nemecek, 1993; Del

Toro et al., 2019; Van Munster, 2020).

Knowledge of the factors affecting transmission efficiency is important in understanding epidemiology and improving the control of viral diseases. The steps in

PVY transmission including acquisition, retention and inoculation (no latent period in

PVY) involve complex and specific molecular interactions that will be briefly discussed in the next paragraphs. Retention times for PVY strains in its vectors range from 4 to

17 hours, however, this also depends on temperature and the aphid species that is feeding (Robert et al., 2000). All PVY strains are transmitted by aphids, yet, the PVYC strain is only transmitted in the presence of a helper component of other PVY strains or PVA (Govier and Kassanis, 1974a, 1974b). PVYC has historically been considered non-aphid-transmissible, yet, a biological study of 8 PVYC isolates suggest that certain isolates are aphid-transmissible (Blanco-Urgoiti et al., 1998). The most important criteria which influence PVY transmission are discussed as below.

Strain specific aspects and virus plant sources

The transmission efficiency studies to measure the transmissibility of PVY strains have

shown controversial results. Results of experiments by Mondal et al. (2016) and

Mondal and Gray (2017) showed that transmissibility of isolates of NTN and N:O strains by M. persicae and R. padi is higher than the ordinary O strain. The NTN strain also showed to be more easily transmitted than O and N:O in experiments by

Srinivasan et al. (2012). Mondal et al. (2016, 2017) pointed to differences in transmissibility of PVY strains as a reason of changes in strain prevalence. The result of these studies showed the recombinant strains NTN and N:O were transmitted more efficient than ordinary O strain, whereas results of recent studies show no significant differences in transmissibility of the N, NTN and N-Wi strains (Verbeek et al.,

2010).The data from the recent experiments by Mondal and Gray (2017), Zsuzsa and

Almasi (2004) and Mello et al. (2011), unlike previous results (Mondal et al., 2016), do not show differences in transmissibility of PVY strains, and they believe that emergence of recombinants (NTN and N:O) in USA potato farms do not relate to the transmission efficiency of PVY strains by M. persicae, however, the transmission efficiency varied when different host potato cultivars were serving as the source of these strains. Higher titers of some strains (NTN) over the other strains in different potato cultivars as a host of virus source could facilitate the transmission efficiency of these strains (Carroll et al., 2016). Differences in the concentration of PVY within the plants, so also between potato cultivars, is another factor explaining the variation in transmission efficiency, as a result of the interaction of virus and host plant (Cervantes and Alvarez, 2011).

Impact of the virus receiving host plants

Host cultivars that are more preferred and hosted by aphids can logically also be more exposed to different strains of PVY. In addition, the plant defense system against PVY and/or the aphid itself, which both may differ between the cultivars of the same plant

species, are key determinants in virus transmission. Symptom development and virus particle concentration for each specific PVY strain can differ in each of the potato cultivars (Dupuis 2017) and this affects transmission and epidemiology of PVY

(Verbeek et al., 2010; Dupuis et al., 2019; Mondal et al., 2017).

Aphid and transmission efficiency

PVY is transmitted by more than 60 aphid species in non-persistent manner (Table

1.5). They transmit PVY with various rate of efficiency from zero to about 83% depending on the experimental conditions such as aphid species (or even biotypes and clones) and virus strain or isolates (Kostiw, 1979; Van Hoof, 1980; Van Harten,

1983; Harrington and Gibson, 1989; Sigvald, 1984, 1992; de Bokx and Piron, 1990;

Halbert et al., 2003; Verbeek et al., 2010; Mondal et al., 2017). Some aphid species such as Aphis fabae, Aulacurthum solani and Acyrthosiphon pisum could not transmit

PVY, or transmitted the virus only in very low rates (Kanavaki et al., 2006; Boquel et al., 2011), whereas Myzus persicae has been reported as the most efficient vector, transmitting PVY with an efficiency of up to 83%. Most of these species do not colonize potato plants and only halt and probe potato leaves briefly (Blackman and Eastop,

2000).

Transmission efficiency of each aphid species that could be determined by the behavioral and physiochemical entity of vectors. Aphid behaviors such as flying, landing, host preference and feeding, among them feeding the most important, are the main behavior that affect the virus transmission efficiency (Lacomme et al., 2017).

Moreover, increasing population densities of efficient aphid vectors on the virus infected plant, increased spread of PVY more significantly (Galimberti et al., 2020).

In 2010, 15 aphid species were recorded in potato crop in Belgium and in 2011 the number of aphid species was higher than in 2010 with 19 species for the same crop

(Vandereycken et al., 2015). In 2010, M. dirhodum, S. avenae, S. fragariae, A. fabae,

A. pisum, M. persicae, R. padi, R. maidis, R. insertum, C. aegopodii, M. rosae, M. euphorbiae, A. solani, Cinara sp. and B. brassicae were collected from the potato crop and in 2011, M. dirhodum, S. fragariae, Aphis fabae, A. craccivora, A. nasturtii, M. viciae, A. pisum, M. persicae, R. padi, R. maidis, C. horni, C. aegopodii, M. rosae, M. euphorbiae, A. solani, Cinara sp., Tetraneura sp., Phyllaphis fagi, P. humuli were collected.

In a four-year survey in Belgium (Wallonia) from 2009 to 2012, about 43 aphid species were detected using yellow pan traps. Although the abundance of aphid species varied from year to year, the most common species encountered were Metopolophium dirhodum (Walker) (28%), Aphis fabae (Scopoli) (18%), Cavariella aegopodii (Scopoli)

(14%) and Myzus persicae (Sulzer) (14%). Four of the five aphid species that normally feed on potatoes (Rolot, 2005; Nanayakkara et al., 2012) were collected: M. persicae

(14%), M. euphorbiae (3%), and A. solani and A. gossypii (less than 1%). Despite intensive sampling attempts, none of them have been found on plants in situ. Although several aphid species were non-colonizers and all species transmitted PVY with differing efficiencies (Ragsdale et al., 2001), only 7% were significant potato virus vectors (Bosquée et al., 2016).

Table 1.5. Aphid species that transmit PVY naturally or experimentally. The aphid species that are known to colonize potato plants are in bold. Adapted from Al Mrabeh et al. (2010)

Transmission No. Aphid species PVY strain (s) References Efficiency (%) DiFonzo et al.,1997; Boquel et al., 2011; 1 Acyrthosiphon pisum PVYO, PVYN 0-14 Kanavaki et al., 2006; Fox et al., 2017 2 Acyrthosiphon primulae PVYN 15 Ragsdale et al., 2001 3 Anoecia corni PVYO - Basky and Raccah, 1990 4 Aphis citricola PVY (pepper) 6.2 Raccah et al.,1985

5 Aphis craccivora PVYO 4 Basky and Raccah, 1990 DiFonzo et al.,1997; Basky and Almasi, PVYO, PVYN, 6 Aphis fabae 0-24 2005; Kanavaki et al., 2006; Boquel et al., PVYNTN 2011; Fox et al., 2017 7 Aphis fabae cirsiacanthoides PVYO, PVYN - Basky and Almasi, 2005 8 Aphis frangulae PVYO 42 Sigvald, 1992 PVYO, PVYN, 9 Aphis glyacines 14 to 75 Davis et al., 2005 PVYNTN 10 Aphis gossypii PVYO 31 Raccah et al., 1985 11 Aphis hellantti PVYO DiFonzo et al., 1997 Sigvald, 1984; Harrington et al., 1986; de 12 Aphis nasturtii PVYO, PVYN 7.1 Bokx and Piron, 1990 van Hoof, 1980; Harrington and Gibson, 13 Aphis pomi PVYO, PVYN 2 to 9 1989; Basky and Almasi, 2005 14 Aphis rumici PVYO - Basky and Raccah 1990 Harrington et al.,1986; Harrington and 15 Aphis sambuci PVYO, PVYN 4.3 to 12 Gibson, 1989; de Bokx and Piron, 1990 16 Aphis spiraecola PVYN - Basky and Almasi 2005 van Hoof 1980; Kanavaki et al., 2006; 17 Aulacorthum solani PVYO, PVYN 0-5 Verbeek et al., 2010; Boquel et al., 2011; 18 Brachycaudus cardui PVY Basky 2002 Edwards, 1963; van Harten, 1983; 19 Brachycaudus helichrysi PVY, PVYO, PVYN 0.9 to12.5 Harrington et al.,1986; de Bokx and Piron, 1990

20 Brevicoryne brassicae PVY, PVYO - Sigvald 1984; Basky and Raccah, 1990

21 Capitophorus elaeagni PVY, PVYO 2 DiFonzo et al.,1997; Halbert et al., 2003

22 Capitophorus hippophaes PVYN 3 van Hoof 1980; de Bokx and Piron, 1990

23 Cavariella aegopodii PVYO, PVYN 0.2 to 0.4 de Bokx and Piron, 1990

24 Cavariella pastinaca PVYN - Salazar, 1996 25 Cryptomyzus ballotae PVYO 100 Harrington et al.,1986 26 Cryptomyzus galeopsidis PVYN 17.4 de Bokx and Piron, 1990 27 Cryptomyzus ribis PVYN 15.4 de Bokx and Piron, 1990 Halbert et al., 2003; Basky and Almasi, 28 Diuraphis noxia PVY, PVYO 4-7 2005 Drepanosiphum PVYN 29 0.6 Powell et al.,1992; Powell et al.,1995 platanoidis 30 Dysaphis plantaginea PVY, PVYO - Basky and Raccah, 1990 de Bokx and Piron, 1990; Harrington and 31 Dysaphis spp PVYN 1.8 Gibson, 1989 32 Hayhurstia atripllicis PVY - Basky and Raccah, 1990 33 Hyadaphis foeniculi PVYN 14.7 de Bokx and Piron, 1990; Piron, 1986 Piron 1986; de Bokx and Piron, 1990; 34 Hyalopterus pruni PVY, PVYO, PVYN 13.9 Basky and Raccah 1990 Harrington et al., 1986; de Bokx and 35 Hyperomyzus lactucae PVYO, PVYN 17.4 Piron, 1990

36 Hyperomyzus pallidus PVY - Basky and Raccah, 1990

37 Lipaphis erysimi PVYO - DiFonzo et al.,1997

van Hoof, 1980; van Harten, 1983; 38 Macrosiphum euphorbiae PVYO, PVYN 29 Harrington et al.,1986; de Bokx and Piron, 1990

39 Macrosiphum rosae PVYN - Basky and Almasi 2005

40 Metopolophium albidum PVYN 11 van Hoof 1980

van Hoof, 1980; van Harten, 1983; de Metopolophium PVY, PVYO, 41 - Bokx and Piron, 1990; Sigvald, 1992; dirhodum PVYN Halbert et al., 2003

42 Metopolophium festucae PVYO 0.5 Harrington et al.,1986

43 Myzaphis rosarum PVYO 10 Harrington et al.,1986

44 Neomyzus circumflexus PVYO, PVYN - Salazar, 1996

PVYN,, 45 Myzus ascaionicus PVYNTN, - Verbeek et al., 2010 PVYNWi Harrington et al.,1986; de Bokx and Piron, 46 Myzus cerasi PVYO, PVYN 3.2 1990; Basky and Almasi, 2005

47 Myzus certus PVYO, PVYN 71 van Hoof, 1980; de Bokx and Piron, 1990

Harrington et al.,1986; Basky and Almasi, 48 Myzus ligustri PVYO, PVYN 50 2005

49 Myzus myosotidis PVYO 100 Harrington et al.,1986

Van Hoof 1980; Piron 1986; Harrington Myzus persicae 50 PVY, PVYN 13.5 and Gibson 1989; Halbert et al., 2003; nicotianae Kanavaki et al., 2006 van Hoof, 1980; van Harten, 1983, Harrington et al.,1986; de Bokx and Piron, PVY, PVYO, 51 Myzus persicae 8.4 to 83 1990; Sigvald 1992; Fereres et al.,1993; PVYN Halbert et al., 2003; Kanavaki et al., 2006; Boquel et al., 2011; Fox et al., 2017 van Hoof, 1980; van Harten, 1983; PVYO, PVYN 52 Phorodon humuli 35 Harrington et al.,1986; de Bokx and Piron,

1990 van Hoof, 1980; van Harten, 1983; 53 Rhopalosiphum insertum PVYO, PVYN 50 Harrington et al.,1986; de Bokx and Piron, 1990 54 Rhopalosiphum maidis PVYO, PVYO 1.5 DiFonzo et al.,1997; Halbert et al., 2003 Kostiw, 1979; van Hoof, 1980; van Harten, 1983; Sigvald, 1984; Harrington PVY, PVYO, et al.,1986; Piron, 1986; Harrington and 55 Rhopalosiphum padi 2 to 11.5 PVYN Gibson, 1989; de Bokx and Piron, 1990; DiFonzo et al.,1997; Halbert et al., 2003; Basky and Almasi, 2005 Rhopalosiphum 56 PVY - Ragsdale et al., 2001 pseudobrassicae 57 Schizaphis graminum PVY, PVYO, - Basky and Raccah, 1990; DiFonzo et

PVYN al.,1997; Halbert et al., 2003; Basky and Almasi, 2005 Sigvald, 1984; Harrington et al.,1986; de Bokx and Piron, 1990; DiFonzo et 58 Sitobion avenae PVYO, PVYN 0.1 to 1.8 al.,1997; Piron, 1986; Harrington and Gibson, 1989 Harrington et al.,1986; Piron 1986; 59 Sitobion fragariae PVYO, PVYN 0.5 to 10.1 Harrington and Gibson, 1989; de Bokx and Piron, 1990 60 Sitobion graminum PVYNTN, PVYNWi - Verbeek et al., 2010 61 Staphylae tulipaellus PVYN - Salazar, 1996 62 Therioaphis trifolli PVY (pepper) - Perez et al.,1995 63 Tetraneura ulmi PVY - Basky and Raccah, 1990 Harrington et al.,1986; Harrington and 64 Uroleucon spp PVYN 0.5 – 8.3 Gibson, 1989 65 Uroleucon sonchi PVY - Raccah et al., 1985

Winged versus wingless aphids

Apterae do not appear to play a significant role in field spread of PVY (Ragsdale et al.,

2001), but both forms of M. persicae transmit PVY. Winged aphids that are more mobile, transmit the virus to further distances (13 m), while wingless aphids hardly walk over distances longer than 1 m. The limited walking distance and low retention times of such NCNP viruses result in a potential effect of wingless aphid types estimated at 2-3% only, in a hypothetical simulation experiment. However the abundance of the wingless form could increase its importance (Ferrar, 1969;

Thygesen, 1968; Ragsdale et al., 1994; Radcliffe and Ragsdale, 2002).

Impact of the feeding behavior

The host plant selection and feeding behavior of the aphid species could be characteristic for each aphid and could affect the virus transmission efficiency. First the color and odors of plants attract aphids to the plants (pre-alighting or before landing). After the landing and assessment of plant surface, they start to briefly probe and puncture the epidermal cell walls to taste sap of plant leaves (this takes a few

seconds) and find a desired host. If aphids find the plant edible, they will stay and find an appropriate site for the next feeding steps, which include saliva secretion (gelling and watery) and extension of the stylet to reach and ingest phloem sap. Otherwise, when they find the plant unpleasant, they stop probing and will leave it to try another plant (Powell et al., 2006; Pettersson et al., 2007; Fereres and Moreno, 2009; Brault et al., 2010). Successful feeding is followed by a complex process that starts with plant selection, landing, probing (insertion of stylets) and phloem sap suction that could be described precisely as: 1) landing behavior, 2) early contacts with plant and surface evaluation before probing, 3) probing the epidermal cells, 4) stylet penetration through mesophyll, 5) sieve element puncturing and phloem salivation and 6) access to phloem and continuing ingestion. However, xylem sap intake is often happen to maintain water balance. (Pompon et al., 2011; Van Emden and Harrington, 2017).

Different feeding behavior could affect transmission efficiency of aphid species.

Boquel et al. (2011) showed that the AAP for M. persicae, M. euphorbiae, A. pisum,

R. padi, S. avenae, B. brassicae and A. fabae, took place in 11, 15, 30, 31,35, 48 and

90 minutes and transmission efficiency of these aphids were 83.3 (a), 26.7 (b), 6.7

(bc), 0 (c), 16.7 (bc), 3.3 (c), 0 (c) %, respectively. Based on the M. persicae and A. fabae data with the shortest and longest AAP, respectively, the results of this study show the relevance of feeding behavior and virus (PVY) transmission. Aphids may refuse feeding in any above-mentioned steps. Primarly the plant color (Doring et al.,

2009; Doring and Chittka, 2007) and odor (Nottingham et al., 1991; Pettersson et al.,

2003) attract the aphids and encourage them to land on (winged), or walk toward

(apterous) the attractive plants. Then they start a brief intracellular probing of epidermal and parenchymal cells as an essential step in non- and semi-persistence transmission of plant viruses (Prado and Tjallingii, 1994; Tjallingii et al., 2010). An

electric penetration graph (EPG) has been used in many different studies to depict the feeding behavior of aphid species and correlation between feeding behavior and virus transmission (Sauvion and Rahbe, 1999; Boquel et al., 2012).

Each of the abovementioned step in feeding behavior is affected by environmental conditions such as temperature, light, humidity, plant volatiles and pesticide application that could change transmission efficiency (Powell et al., 2006; Fereres and

Moreno, 2009; Boquel et al., 2014).

Environmental conditions

Effects of environmental conditions such as temperature, relative humidity, light (Singh et al., 1988), wind (Kumar et al., 2017) and CO2 concentration on virus transmission have, to some extent, been studied. These environmental components could change vector behavior, plant growth or reactions to the virus and virus replication in the host plants, and therefore transmission efficiency.

Aphid behavior in general and feeding behavior of aphids in particular, are affected by different environmental conditions too, including: wind, light and photoperiod, temperature, atmospheric humidity, plant drought, colors, volatiles of plants, starving and plants or diet that the aphids ate before starting new feeding, source plants of the virus and receptor plant.

Favored temperatures increases aphid reproduction rate in general (Lamb and White,

1966), and as a consequence, the risk of virus transmission will increase. Singh et al.

(1988) showed that temperature and relative humidity (RH) affect transmission of both

PVY and PLRV viruses by 30%–35%, while light intensity did not. The effect of temperature on inoculated plant is also a determinant in transmission and virus proliferation in plants (Chung et al., 2016). The Ny genes in resistance cultivars to PVY strains, can be overcome at a temperature 28ºC, however the symptoms range from

cryptic to mosaic and leaf drop (Valkonen, 1997; Qian et al., 2016; Makarova et al.,

2018).

Drought elicits plant hormone secretion and volatile emission. As reviewed by

Szczepaniec and Finke (2019) and van Munster (2020), stressed plants may put a positive, negative or neutral impact on aphid behaviors as common vector of NCNP viruses.

Transmission efficiency is affected by CO2 concentration however, 2 different non- persistent virus systems (PVY and CMV), showed opposite results. PVY transmission rate increased whereas CMV decreased under similar different CO2 treatments. The effect was shown to originate from the plant physiological variation, not from aphid behavior alteration (Ye et al., 2010; Dader et al., 2016; Bosquee et al., 2018). In another study, a combination of increased temperature and CO2 levels decreased

PVY concentration in tobacco plants that lead to reduced PVY transmission by M. persicae (Del Toro et al., 2019).

Meteorological parameters play a crucial role in this migration of aphids and wind helps aphids to migrate. They could surf on wind and spread virus in a longer distance as the aphid flight and wind direction showed high correlation (Ghosh et al., 2019).

Moreover, wind affect flying behavior, as wind speeds above 2.4 km/hr. can delay and reduce takeoff of most winged aphids (Kring, 1972; Walters and Dixon, 1984). Seed potato production in high altitude area is recommended due to low temperature and wind blowing limiting aphid activities (Johnson and Powelson 2008). Rainfall as another climate factor has negative effect on aphid populations and PVY prevalence, however it depends on time and type of the rainfall and crop growth stage (Kumar et al., 2017).

Plant volatiles, color and nutrients

Aphids select plants to feed, first by color, odors of plant volatiles and then nutrient composition. Aphids respond to colors for distances of about one meter (Kring, 1972), and move toward or land to access the favorite plants. Color influences the attraction of aphids to plants. During the flight, they respond to visual stimuli (Kring, 1972) and discriminate green color of host plants from the soil background (Kennedy et al., 1962).

Doring and Chittka (2007) demonstrate that the aphid behaviors for some species correlated positive as well as negative to the green and blue and/ or UV color ranges, respectively. They also showed that attraction to the color yellow is relevant to its brightness effects.

Potyviruses and indeed PVY can influence host plant characteristics such as volatile emissions. The EPG technique reveals that depending on the colonizing (Myzus persicae) and non-colonizing (transitory) (Aphis fabae, Brevicoryne brassicae, and

Sitobion avenae) aphids, PVY-infected plants modify the aphid behavior and PVY transmission possibility (Boquel et al., 2012; Gadhave et al., 2019). Results of Bak et al. (2019) indicate induction of ethylene signaling by PVY, attracts aphids to infected plants and mediated virus spread. Ahmed et al. (2019) showed that the wingless M. persicae were efﬁcient to distinguish plant cultivar differences by leaf nitrogen content

(Ahmed et al., 2019).

Virus infected plants mediate vector orientation, feeding, and dispersal by changing color or emissions (Mauck et al., 2010; Mauck, 2016). In addition, the transmission activation (TA) phenomenon in infected plants of two transmissible NCNP and NCSP viruses (TuMV; Potyviridae and CaMV; Caulimoviridae) has recently been reported, where the virus produces transmissible complexes that are effectively acquired and transmitted in the presence of vectors in the host plant (Berthelot et al., 2019).

Control measures; Insecticides, straw mulch and mineral oil

Pesticides change vector behavior and can lead to lower or higher PVY transmission rates by increased mortality, and decreased probing or by increased flights and walking around. Insecticides and antagonists, which can reduce the incidence of the persistently transmitted PLRV, are hardly effective against the non-persistently transmitted PVY, because they do not kill the vectors fast enough to prevent transmission (Shanks and Chapman, 1965; Boquel et al., 2014).

Straw mulch reduced the incidence of PVY significantly in the progeny tubers, with the reduction ranging from 50% to 70% in all three years. The straw mulching could impact aphid sight and reduces the contrast between the plant background canopy (Doring and Schmidt, 2007; Doring, 2014). Application of mineral oil reduced the time of PVY retention in the stylet of M. persicae from 17 h to just 2 min (Wrobel, 2009). The incidence of PVY was reduced by 43% to 58% with mineral oil over two years and by

25% over one year with the synthetic pyrethroid insecticide esfenvalerate (Kirchner et al., 2014).

Mineral oils as a low cost and low toxic material have been reported to reduce the transmission of non-persistent aphid-borne viruses by changing feeding behavior and colonization of aphids (Ameline et al., 2009; Ameline et al., 2010; Ouyang et al., 2013), killing aphids fast (Galimberti and Alyokhin, 2018), and/or decreasing virus retention in the aphid stylet by covering the inner side of aphid stylet and preventing virus particles acquisition (Powell and Hardie, 1994; Wang and Pirone, 1996; Boquel et al.,

2013).

Virus load

Feeding on the plants as a source of virus and also plants that receive the virus from aphid vectors are the two important sources of variation that affect directly virus

transmission efficiency by vectors. The load of the virus in source plant and taste and acceptability of source and recipient plants for aphid vector affect transmission directly

(Pirone and Thornburym, 1988; Verbeek et al., 2010; Fox et al., 2017). However, aphid host plants could also influence plant virus transmission indirectly.

Vector plant hosts before virus acquisition

The plant species on which the aphids were reared is also an important variable in the virus transmission, yet one which has not been studied extensively. Many research has focused on plant species and application of their metabolite (essential oils, flavonoids, lectins, alkaloids, glycosides, esters and fatty acids) to combat insect pests as repellent, toxicants, growth retardants, chemosterilants, and attractants (Loris,

2002; reviewed in Hikal et al., 2017). The effect of some plant compounds, such as extracted oil and lectins, are being studied for their effects on plant virus transmission by insect vectors; but effects of alive plant as a source of these components is not widely studied.

A preliminary study on three different plant species for rearing of M. persicae (potato, tobacco and oilseed rape) revealed a lower transmission efficiency of some PVY isolates by aphids reared on potato (Al-Mrabeh, 2010). These transmission differences might be linked to the composition of the host plant on which the aphids were reared. In addition, some plant species like broad bean (Vicia faba L.) are a source of lectins (Loewus and Tanner, 1982), of which some have proven to interfere with the transmission of PVY and cucumber mosaic virus (CMV) (Bosquee et al.,

2014). Additionally, Killiny et al. (2011) confirmed a significant effect of two plant lectins

(wheat germ agglutinin, concanavalin A) in disrupting the vector transmission of Xylella fastidiosa.

Recent studies showed that PVY transmission were reduced by feeding aphid vectors

on extracted lectins of pea and Broad bean (Francis et al., 2020). Moreover, they showed that the transmission of Barley yellow dwarf and Pea enation mosaic virus

(both CPNP virus) was also reduced when aphids were reared on artificial diet containing the lectin Concanavalin A. Lectins are proteins that bind to specific carbohydrate structures and could be potentiality a competitor in plant virus transmission by aphids, and even in human virus diseases such as human immunodeficiency virus (HIV) and coronaviruses (Balzarini et al., 2004; Killiny et al.,

2011; Mitchell et al., 2017). These lectins could potentially reduce plant transmission by binding to receptors located in the stylet or gut of the insect (Naidu et al., 2004) or viral glycoproteins (Tang et al., 2015). Moreover, the aphid feeding behavior (the number and duration of aphid probes) significantly reduced when they fed on artificial diet containing lectins (Sprawka and Goławska, 2009). Feeding behavior and virus transmission are correlated, as discussed.

PLRV transmission efficiency significantly reduced by turnip-reared aphids compared to Physalis-reared aphids in a non lectin content plants. Proteome analysis revealed that lysosomal enzymes and other cysteine proteases (cathB) were increased in turnip-reared aphids and indirectly decreased virus transmission (Pinheiro et al.,

2017). Symmes and Perring (2007) also showed transmission of ZYMV (another NP transmissible potyvirus) by M. persicae was affected by the host plant on which the aphid is reared as well as the host plant on which it feeds just before virus acquisition

(short term preacquisition). A few Clones of M. persicae collected from different hosts

(Malva parviflora L. and Brassica oleracea L.) and/or the same clone reared and/or shortly fed on the different host (Abelmoschus esculentus L. and Brassica juncea L.) showed significant differences in ZYMV transmission (Symmes and Perring, 2007).

Starving, acquisition accession period (AAP), inoculation accession period

(IAP) and virus retention

Starving of aphids before non-persistent virus acquisition increases virus transmission

(Taylor and Robertson, 1974; Powell, 1993) by changing feeding habit (probing behavior) and increasing the appetite of aphids. Starving by itself, does not universally increase the transmission rate of non-circulative viruses (Jimenez et al., 2017). In many plant virus transmission studies, aphids are left to starve for 2-3 h at the start in order to be sure about aphid feeding on the infected plants (acquisition) and later the healthy test plants (inoculation). The AAP for NCNP viruses is very short (1-5 min) and extending this period leads to a decreased virus transmission efficiency (Al-Mrabeh,

2010; Fereres and Moreno, 2009; Watson and Roberts, 1939). Kotzampigikis et al.

(2009) showed that the optimal period for acquisition (AAP) and inoculation (IAP) of

PVY by the aphids is 30 s and the minimal is 1 s. They found that M. persicae remains viruliferous from 2 to 4 h and depends on the number of visited healthy plants and feeding time on them, as the retention period in aphid vectors decreases by the number and time of feeding.

The non-persistent plant viruses show a low level of vector speciﬁcity because they are transmitted by many aphid species (Pirone and Harris, 1977). Feeding behaviors or host plant selection and properties of virus receptor of each aphid species

(compatibility with the helper component and/or availability of receptor) (Uzest et al.,

2007) could be determinants in virus transmission efficiency attributed to aphids

(Nanayakkara et al., 2012b). These parameters affect aphid feeding behavior and lead to a change in the transmission efficiency in turn and the virus epidemiology. But the potential transmission efficiency of individual aphids in optimal conditions is determined by molecular properties of aphids.

Molecular aspects of PVY transmission by aphids

PVY acquisition and inoculation as an essential part of virus transmission take place during aphid’s short probe (less than a minute) of plant epidermal cells to evaluate plant edibility. Helper component (HCpro) and coat protein (CP) of potyviruses, and receptor molecules of aphids are the three important proteins which are involved in virus transmission by aphids (Nault and Styer, 1972; Powell, 2005).

Electric penetration graph (EPG), electron microscopy (EM) imaging and aphid transmission bioassays have shown that acquisition occurred mainly during the last sub-phase II (II-3) and the virus particles in the common duct (CD) are inoculated to plants by salivation in first sub-phase II during next probes (Fig. 1.18) (Ammar et al.,

1994; Fereres and Collar, 2001). This knowledge led to propose ingestion-salivation hypotheses that is more accepted rather than other two hypotheses (ingestion- egestion and conformational change) (Ammar et al., 1994; Salomon and Bernardi,

1995; Pirone and Blanc, 1996; Martin et al., 1997; Fereres and Collar, 2001; Pirone and Perry 2002; Powell, 2005; Fereres, 2007).

Figure 1.18. Intracellular sub-phases (II-1, II-2 and II-3) correlated with non- persistent virus inoculation (bottom left) and acquisition (bottom right). Watery saliva send trapped virus particles into the plant protoplast from CD (common duct) (Adapted from: Martin et al.,1997).

HCPro is a multifunctional protein (listed in Table 1.3) (52 KDa, 456 AA and 285-740 genome) (Kerlan, 2006; Valli et al., 2018) that was proposed to join virus particles and putative receptors in stylet as a bridge (Bridge hypothesis) (Pirone and Blanc, 1996)

(Fig. 1.18) to transmit potyviruses. Two domains of the HC of PVY, including Pro-Thr-

Lys (PTK) that interact with Asp-Ala-Gly (DAG) motif of CP subunits in the virus particle (conserved in potyviruses and PVY strains) (Kerlan 2006) and the Lys-Ile-Thr-

Cys (KITC) that could be involved in CuP interaction in aphid, are essential in virus transmission by aphids (Thornbury et al.,1990). A change in the amino acids of either motives will result in loss of helper function, however the sequences of the KITC in potyviruses may be substituted by others, while the function remains similar in other viruses; KITC in PVY, TVMV and TEV (Thornbury et al.,1990), KLSC in ZYMV

(Granier et al., 1993) soybean mosaic virus (SBMV) (Jayaram et al., 1992); bean common mosaic virus (Fang et al.,1995) and peanut stripe virus (Flasinski and

Cassidi, 1998), QITC in yam mosaic virus (Aleman et al., 1996); RTTC in sweet potato feathery mottle virus (Sakai et al.,1977); KLTC in pepper mottle virus (Vance et al.,

1992); RITC in clover yellow vein virus (Takahashi et al.,1997), bean yellow mosaic virus (BYMV) (Guyatt et al., 1996), pea seed-borne mosaic virus (Johansen et al.,

1991) and KVSC in peanut mottle virus (Flasinski and Cassidi, 1998). These variations in the binding site of the HC to CuP of aphid could be one of the reasons for aphid specificity in Potyvirus transmission (Fig. 1.19). Physical and chemical properties and the location of receptor/s in aphids are a critical aspects in PVY transmission, however it is not well characterized compare to the other two necessary component (CP and

HCpro) in transmission. Early researchers reported that the non-persistently transmitted PVY was associated with the cuticular distal parts of the aphids’ stylets

(Bradley and Ganong, 1955a, b) and which was later confirmed using electron

microscopy imaging (Taylor and Robertson, 1974; Ammar et al., 1994). However, there was no evidence to support virus transmission. Later, a combination of aphid transmission tests by functional and nonfunctional CP and HC of Potyvirus and stylet

Immunogold labeling electron microscopy showed that loss of aphid transmissibility of mutant (HC and/or CP) potyviruses is due to the non-retention virions in the mouthparts (Wang et al. 1996, 1998). These findings supported the proposed concept about involvement of the retained Potyvirus particles in aphid transmission.

Fig. 1.19. Bridge hypothesis in aphid transmission of PVY. Left: an aphid is feeding from an infected plant. Centre: longitudinal section of the mandibular stylet Right: a helper component proteinase (HCPro) complex.

In a different virus helper component system but similar (semi-persistent virus transmission system with P2 protein instead of HCpro), Uzest et al. (2007, 2010) showed localization of the Cauliflower mosaic virus (CaMV) and putative aphid receptors on the tip of the stylet of Acyrthosiphon pisum, M. persicae, and Brevicoryne brassicae, but not in a non-vectoring species, using green fluorescent protein (GFP) labeling. They found that P2 protein bind to “acrostyle” (a new structure on the tip of the stylet in common duct), that is very similar in terms of location, shape, size, and

perhaps biochemical composition in other aphids. They also showed the receptors of

CaMV in some non-vector aphids immerse in cuticle and are inaccessible to bind the

P2. In vitro interaction assay between HC and extracted or expressed proteins of aphids have showed that Cuticlar Proteins (CuP) are the most probable molecules as receptors of semi and non-persistent transmission viruses (Dombrovsky et al., 2007a, b; Uzest et al., 2007, 2010; Al-Mrabeh, 2010). Al-Mrabeh (2010) and Fernandez-

Calvino et al. (2010) revealed some extra interactions between TEV HCpro and ribosomal protein S2 (RPS2), exoskeleton protein, beta-tubulin, ATP citrate lyase, serine/threonine-protein phosphatase, and a membrane protein. These protein interactions seem to be unrelated to the aphid transmission, and may be due to the multifunctional properties of the HCpro or an unspecific interaction.

Based on the above mentioned findings, CuPs are the most likely receptors of potyviruses and CaMV in the tip of aphid’s stylet. The largest structural CuP groups in arthropods, the CPR family, is classified base on a 28-aa conserved motif named the

“R&R Consensus” (Rebers and Riddiford, 1988) that binds to chitin (Rebers and Willis,

2001; Togawa et al., 2008), which may enhance the physical properties of the cuticle.

RR-1 CuPs have been isolated from soft or flexible cuticles, whereas RR-2 CuPs more often associated with hard cuticles (discussed by Willis et al., 2005). Most CuP genes are regulated by moulting hormones such as juvenile hormones and ecdysteroids

(Riddiford et al., 2003). A recent study showed the vital role of mpcp1 CuP in M. persicae, as when its gene was knocked down, fecundity of the aphid reduced by 40 to 47% (Bhatia and Bhattacharya, 2018). Additionally, the expression of some CuPs

(MPCP1, MPCP2, MPCP4, MPCP5) in M. persicae increased following feeding on

CMV-infected plants (Liang and Gao, 2017).

The RR sub-groups genes of CPR family belonged to multi-gene families and are often

clustered in the genome and co-regulated (Cornman et al., 2008; Togawa et al., 2008).

CuP genes could be regulated at the adult stage, when no moulting process occurs anymore. Cuticle deposition can be controlled by a circadian clock in many insects (Ito et al., 2008). Gallot et al. (2010) demonstrated that genes encoding CPs and genes involved in cuticle modification processes are regulated by photoperiod shortening.

As mentioned, receptor molecules of CaMV are submersed in a proteinaceous structure at stylet’s tip (Uzest et al., 2007). They later demonstrated specific antisera raised against a peptide termed pepL (GSYSLLEADGSTRTVE) not only binds to the acrostylet in the CD but also binds to P2 protein of CaMV. The pepL represents the

RR2 conserved motif in 20 CuPs of A. pisum (Uzest et al., 2010). On the other hand, the HC of an NCNP Potyvirus (ZYMV), interacted with nine different CuPs extracted from M. persicae, but not with expressed CuPs in a cDNA library (Mpcp1, Mpcp2,

Mpcp3, Mpcp4, Mpcp5), maybe due to non-functional expression. Extracted proteins

3, 4, 5 and 9 represent peptides that were present in aphid CuPs that were deposited in GenBank. Peptides in these four proteins included a conserved amino acid sequence found in the R&R consensus (Dombrovsky et al., 2007b).

Afterwards, the roles of the RR1 and RR2 CuPs groups were determined in HC and

P2 protein interactions (Dombrovsky et al., 2007a, b; Uzest et al., 2010). More data on involvement of CuP in a CPP transmission system were provided when they showed interacting between the CPR1 CuP of the plant hopper Laodelphax striatellus and nucleocapsid protein (pC3) of rice stripe tenuivirus (RSV) by yeast-2-hybrid approach and silencing CPR1 (Liu et al., 2015).

Webster et al. (2017) revealed that the acrostylet is a complex structure contains CuPs

(mainly from the RR-2 family) with available and/or immersed domains potentially involved in virus-insect interactions. Even so they showed accessibility of different

regions but a few amino acids of the RR-2 domains on the CuP surface of the acrostylet that one of those interacted with CMV and hence could be accessible at the surface of the acrostyle, however receptor of CaMV did not interacted P2 protein unless treated with chitinase in some aphids. The chitinase treatment increased the accessibility of the immersed domain of CuP and provides better interaction with RR2- antibodies, indicating submerging states of RR2 CuPs in cuticle of acrostylet. Later

Webster et al. (2018) localized two RR1 CuPs (Stylin-01 and Stylin-02) in surface of the acrostylet of both M. persicae that interacted with the CaMV-P2 binding sites and they confirmed that Stylin-01 involved in CaMV transmission using the RNAi technique. The results of an analysis of the proteomes of stylet and 3 cuticular anatomical structures and localization of the CuPs of the pea aphid A. pisum by

Deshoux et al. (2020), indicated that RR1 CuP in the surface of the inner layer of stylet are the most frequent and prime receptor candidate for NC virus (CaMV), whereas

RR2 CuPs are immersed and inaccessible in cuticle for viruses. They could not find any ortholog of MPCP2, (an RR2 CuP) in the MS proteome of leg, wing, antenna and stylet and concluded RR2 CuPs are not an important receptor candidate for NCNP transmissible viruses. However, they did not ignore their role based on several reports

(Domrovski et al., 2007a; Uzest et al., 2007). Moreover Giordano et al. (2020) recently found the ortholog of MPCP2, a RR-2 protein in A. glycines (referred as AG6024500).

The retention of CMV was decreased when the gene encoding MPCP4 (RR1)

(DQ108938) was silenced by 48 h of feeding on dsRNA. Additionally, the expression of some CuPs (MPCP1, MPCP2, MPCP4, MPCP5) in M. persicae increased following feeding on CMV-infected plants (Liang and Gao, 2017).

1.4. Techniques to study virus transmission

To demonstrate a plant virus is transmitted by a given vector, a variety of methods are

available, depending on virus, vector, transmission modes and available instruments.

1.4.1. Transmission bioassays

In general, vector transmission bioassays have been carried out to study vector specificity, identification vector and its efficiency, vector transmission control for different plant viruses. Bioassays are the oldest method to study viruses providing valuable information about epidemiology and control strategy (Dijkstra and Jager,

1998). Feeding aphids (and/or other sucking insects) on artificial diet through parafilm is a widely used bioassay in studies relevant to virus transmission. Depending on the model of virus transmission (Table 1.6), different types of bioassays are carried out.

The tests are mainly designed based on the vector and the type of virus transmission

(Non-circulative NP, NCSP, CNP, and PCP. Rearing and preparing a clone of vectors, providing virus sources (infected plant) and test plants are the first step. The required time for each steps in transmission “Acquisition period, latent period and Inoculation period” depends on type of transmission that are summarized as presented below

(Bragard et al., 2013).

Table 1.6. The most important characteristics of virus transmission types. (s: seconds, m: minutes, h: hours, d: days) (Adapted from: Bragard et al., 2013)

Transmission (Non- Non-circulative Persistent: Persistent: type circulative non- Semi- Circulative, non- Circulative, persistent (NP) persistent propagative (CNP) propagative (CP) Characters (NCSP) Acquisition s-m m h-d h-d period latent period - - + + Retention time m-h m-h h-d/life d-life Inoculation period s-m s-h h-d h-life Virus example Potyviruses Closterovirus Luteovirus Rhabdovirus Vectors example Aphids Aphids Aphids Planthoppers

1.4.2. Electric penetration graph

The electrical penetration graph (EPG) is a method used by biologists to study relationships of insect (aphids, thrips and leafhoppers) with plants and was developed and modified by Mclean and Kinsey (1964) and Tjallingii (1978, 1988). The EPG system uses the insect (sucking insects) and the plants as the components of an electrical circuit and record electric current when the insects start feeding (Fig. 1.20).

The EPG signal displays three distinct sub-phases during intracellular salivation, including II-1, II-2 and II-3 which are associated with the acquisition (II-3), and inoculation (II-1) of the NCNP viruses (Martin et al., 1997; Powell, 2005).

Fig. 1.20. EPG sub-phases II-1 and II-3 have been associated to the inoculation and acquisition of typical non-persistent viruses, respectively (Adapted from: Martin et al., 1997).

This (EPG) method was used to measure specific AAP for aphid species and clarify the transmission steps (Powell 2005; Boquel et al., 2011). Baquel et al. (2011) determined the time between placed aphids on the plant and first probe and AAP for few aphids and PVY using the EPG instrument. Moreover, an EPG designed test by

Powell et al. (1995) and Martin et al. (1997) demonstrated that acquisition and

inoculation of NCNP viruses is associated with the first intracellular probing during sub-phases II-3 and II-1 respectively. Martin et al. (1997) proposed the ingestion- salivation hypothesis as an alternative mechanism to the ingestion–egestion hypothesis on basis EPG tests. In general, EPG technology has been used in insect and insect vector behavior studies (mainly sucking insects) as a valuable method for assessing the importance of behavioral change, virus transmission and/or environmental determinants.

1.4.3. Electron microscopy (EM) and labeling

Direct visualization of pathogens in situ allows researchers to find out the exact replication site in plant cells, changes in the infected cell, and the virus entity.

Transmission Electron Microscopy (TEM) also opened up new horizon for the detection and diagnosis of plant viruses since TMV was pictured in 1930-1939, and the detailed structures of the viruses along with the EM technological development have been established (in: Hull, 2014; Richert-Poggeler et al., 2019) simultaneously.

In addition to virus morphology, virus-vector relations were figured out using in situ

EM, Ultramicrotomy, and different labeling methods including immune and immunogold labeling. Earlier efforts to detect PVY in aphids failed (Kikumoto and

Matsui, 1962), however later not only PVY and other NCNP potyviruses were detected in stylet of aphids (Ammar et al., 1994; Wang et al., 1996), but also the bridge hypothesis and the importance of HC were explained. Recently, a new ultrastructure in the common duct of aphids named acrostylet and its importance for NCSP transmissible CaMV (Caulimovirus) were described by TEM and immunolabelling

(Uzest et al., 2007; Uzest et al., 2010; Webster et al., 2018).

1.4.4. Immunological methods

Virus transmission studies and detection of plant viruses in the body of the vectors is

an important part of plant virus epidemiology. However, presence of the virus in the body of the vectors is not an indication of virus transmissibility. Immunological identification methods as a sensitive detection methods were the most common detection methods before the invention of molecular methods, and is still used as the method of choice in routine virus detection tests. The enzyme-linked immunosorbent assay (ELISA) (Clark and Adams, 1977) as one of the most sensitive serological methods has been used in many studies to detect plant viruses, amongst PVY, in the body of vectors since 1978, starting with CMV (Gera et al., 1978; Carlebach et al.,

1982; Varveri, 2000). Carlebach et al. (1982) claimed PVY detection using ELISA in a single crushed aphid, but the low differences in the OD of viruferous and virus-free aphid suggested that this method still needed to be improved for a reliable virus detection in single aphid. Evaluation of the ELISA test in virus detection in aphids compared to molecular methods, reveals that IC-PCR offered a 1000-fold sensitive detection than ELISA also PCR-ELISA 100-fold than IC-PCR (Varveri, 2000). Higher sensitivity of molecular methods, as well as limitations in virus detection in aphis by

ELISA and the fact that virus detection in the body is not a reliable index for whether or not the insect is a vector, lead to a shift towards molecular methods for virus detection in vectors, although the combination of immunology and labeling is still important for virus localization in vector body.

1.4.5. Molecular methods (q(RT-)PCR and (RT-)PCR)

Different molecular methods are used to detect plant viruses including thermal cycling amplification (Polymerase chain reaction (PCR), Multiplex PCR, Nested PCR, Real- time PCR) and isothermal amplification based methods (Loop-mediated isothermal amplification (LAMP) and Nucleic acids sequence-based amplification (NASBA)). In addition, different methods of genome sequencing are available in order to identify

and differentiated the amplified genome target. Next generation sequencing (NGS) as one of the latest genome sequencing technology provide a wide range opportunity to detect, accurate identification, quantification, and study the ecology, epidemiology, replication and transcription of viruses, viroids and microbiomes (Cao et al., 2017;

Rubio et al., 2020).

PCR as a common laboratory technique makes numerous copies of DNA or complimentary RNA (after revers transcription) templates by 25-40 cycles of thermal treatment to denature double stranded nucleic acid, annealing the primer to desired motives and extending the new strand of nucleic acid. This process take place in the presence of primers, thermo stable DNA polymerase, nucleotides dNTPs and

PCR buffers. PCR has been improving since it was invented by Mullis (1985), while it was initiated when Kleppe (1971) described a process for replicating nucleic acid in the tube (Mullis et al., 1986). PCR and its variant have been used as a highly accurate and sensitive identification tool in plant virology to identify viruses in plants in addition to the body of the vectors.

PCR application for the accurate detection of plant viruses in their vectors provide useful information about virus epidemiology and therefore lead to new virus management strategy.

Successful application of conventional RT-PCR techniques for the detection of plant viruses in their vectors dates to 1992 when Lopez-Moya and his colleague detected two isolates of CaMV (transmissible and non-transmissible), a NCSP transmissible plant virus, in aphid vectors. This is a significant move, but both isolates were detected in aphids, and this method did not reveal anything about the transmission of viruses.

Later, Singh et al. (1996) developed a protocol to detect PVY in a single aphid. After that, Varveri (2000) used Immunocapture PCR (IC-PCR) and print-capture (PC-PCR),

Cambra et al. (2004) detected PPV by nested PCR and Olmos et al. (2005) found that quantity of PPV in complete body of aphids are 40 to more than 2 × 103 particle by real time RT-PCR. Moreno et al. (2007) observed lettuce mosaic virus (LMV) in single vector and non-vector aphids by (IC)-RT-nested-PCR and found no relevance between virus identification in aphids and transmitting capacity, as shown by Lopez-

Moya et al. (1992) for CaMV.

Detection of PVY in stylet as an effort to find a lab index for virus transmission capacity of vectors was done by Zhang et al. (2013). They detected PVY in the 38% of individual stylets of M. persicae that is similar to PVY transmission by M. persicae, however the data of transmission efficiency originated from another study by Moreno et al. (2007).

Boquel et al. (2013) used PVY detection in stylet by RT-PCR to determine the effect of mineral oil in virus acquisition and showed virus decrease in stylet of treated aphids.

Even so, there was no connection between virulifer aphids (virus in stylus) and vector capability using a bioassay transmission. Kim et al. (2016) again found PVY in the entire body of a single aphid by RT-PCR facilitated boiling technique. PVY was quantified in the single detached stylets by (Khelifa, 2019) real time RT-PCR and PVY target copies in positive stylets ranged from 1,141 to 1,904, but about one sample that reach 3,327 copies. This study coincided with our study, in which we investigated the relationship between transmission efficiency and the virus quantity at the aphid stylet

(Bahrami Kamangar, 2019b). As the common duct is about 1% of the stylet length

(Forbes, 1969) it is also the effective site for transmission of NCNP viruses.

1.4.6. Yeast two hybrid system

Protein-protein interactions are essential for all cellular processes, such as DNA replication, transcription, translation, secretion, cell cycle control, metabolism, cell structure development, and enzyme assemblies. In short, the yeast two-hybrid system

is an in vivo technique (in yeast cells) to determine protein-protein interactions. In this system the reporter genes get expressed when the proteins which are under investigation interact. A transcription factor (TF) that bind to DNA and recruit RNA polymerase II contains two domain; the DNA Binding Domain (DBD) which bind the gene promoter on DNA, and an activator Domain (AD) that recruit the RNA polymerase II. These separated parts are fused with the two investigated proteins and expressed using two separate plasmids in yeast cell lines. If the domains of these two proteins could interact then DBD and AD fuse together to rebuilt the TF. This in turn connect to promoter and recruits RNA polymerase II to express the reporter gene (in:

Bruckner et al., 2009). This technic has been applied to evaluate the bridge hypothesis and the HCPro function in potyviruses transmission (Urcuqui-Inchima et al., 1999; Guo et al.,1999; Guo et al., 2001; Kang et al., 2004) and in CMV and rice stripe tenuivirus

(RSV) as well (Liu et al., 2015; Liang and Gao, 2017).

1.4.7. RNAi (RNA interference)

Gene expression can be regulated during both transcription and/or translation as a natural mechanism in living organisms. RNA interference (RNAi) or Post-

Transcriptional Gene Silencing (PTGS) is a natural sequence-specific gene silencing system. RNAi induces the suppression of exogenous (to defend against alien) or endogenous (gene expression regulation) nucleic acids in most eukaryotes and as a defense system, it can degrade viral RNA or mRNA (in DNA viruses) in animals

(Haasnoot et al., 2003; Bronkhorst et al., 2013) and plants (Vance and Vaucheret,

2001). Through an evolutionary interaction between host plant RNAi and target virues, some viruses have developed ways to counter the plant RNAi defense system. For example, the multifunctional HCPro, which is involved in aphid transmission of potyviruses can also suppress the plant RNAi defense systems (Lewsey et al., 2009;

Rodamilans et al., 2018; Pollari et al., 2020).

RNAi as a laboratory technique was first invented to manipulate gene expression in nematode species (Caenorhabditis elegans) by Andrew Z. Fire and Craig C. Mello

(Fire et al., 1998), who were awarded 2006 Nobel Prize in Physiology or Medicine for this discovery. RNAi has since then been used as a powerful laboratory tool to study biological processes and gene functions in different organisms. Moreover, it is a promising tool in both medical field and agriculture for gene therapy and pest management, respectively (Agrawal et al., 2003; Huvenne and Smagghe, 2010; Zhu,

2013; Setten et al., 2019; Wesley, 2019). The RNAi mechanism is induced by a variety of small non coding RNAs:

 micro RNAs (miRNA), about 22 nt endogenous RNA with a unique secondary

structure (Bartel, 2009).

 small interfering or short interfering RNAs (siRNA), about 20-25 base pairs in

length (lam et al., 2015).

 Piwi-interacting RNAs (piRNA), about 21-35 nucleotides and is involved in the

silencing of transposable elements and the regulation of other genetic elements

in germ line cells of animals (Seto et al., 2007).

 Short hairpin RNAs (shRNA) (is an artificial RNA molecule) (Paddison et al.,

2002).

 Paperclip RNAs (pcRNAs) is a synthetic dsRNA with a paperclip structure, (with

two closed ends but not a covalently sealed structure), did not rely on the

conventional, clathrin-mediated uptake processes to facilitate RNAi (Abbasi et

al., 2020).

Both miRNAs and siRNAs are about 22 nt long, but siRNAs are dsRNAs with a

defensive role that originated from alien nucleic acids such as viruses, whereas

miRNAs as regulators of endogenous genes are transcript derived ssRNAs that

fold back on themselves and form a stem loop or hairpin-like shape. The siRNA is

more specific in gene silencing while miRNA is not, and as such a miRNA could

silence several genes partially. RNAi as a molecular tool relies on the production

and delivery of dsRNA to target gene expression in organisms (Fig. 1.21.) (Aalto

et al., 2007; Carthew and Sontheimer, 2009; Lam et al., 2015; Kanakala and

Ghanim, 2016). Below is a brief description of the “siRNA pathway”, which is of

relevance to this thesis and will be henceforth simply refered to as “RNAi”:

 The RNAi pathway is triggered by delivery of dsRNA.

 When dsRNA strands are detected in a cell, the RNase III ribonuclease Dicer

(DCR-2) cleaves it into small interfering RNA (siRNA, 19–24 nucleotides)

 The siRNAs then associate with an enzyme complex to form the RNAi-induced

silencing complex RISC (containing Argonaute proteins).

 RISC undergoes conformational changes which ultimately results in the lead

strand of the siRNA within the complex to recognize and base pair with the

complementary region on the target mRNA sequence.

 This complementary sequence-specific base pairing leads to either the

degradation or the repression of the translation of the target mRNA.

A few studies have focused on RNAi to identify receptors of NCNP aphid-transmitted plant virus including potyviruses, cucumoviruses and Caulimovirus (Liu et al., 2015;

Liang and Gao, 2017; Webster et al., 2017, 2018; Deshoux et al., 2018, 2020), since this method was initiated by Matzke et al. (1989). RNAi-based silencing of CuP genes has been exploited for both pest control (Shang et al., 2019) and for physiological studies, such as in metamorphosis (Jan et al., 2017). RNAi has also been exploited to

control plant virus transmission by virus replication suppression in its vector (Ying et al., 2020).

DsRNA delivery through microinjection, feeding or soaking have all been used as methods to ensure delivery of dsRNA to insects in RNAi research. Improved delivery through feeding employing synthetic nanoparticles to protect/deliver dsRNA or engineered microorganisms that generate dsRNA have been demonstrated to improve the effectiveness of RNAi in insects (Kolliopoulou et al., 2017; Yu et al., 2017;

Christiaens et al., 2020).

RNAi, a promising new approach in gene function research and pest management, is hampered by inefficiencies that limit its use. When using various insect species, strains, developmental stages, tissues, and target genes, the results can be very variable. These shortcomings make widespread use of RNAi-based pest control techniques difficult. Experiments to determine RNAi mechanism have shown that double-stranded ribonucleases (dsRNases), endosomal entrapment, a lack of core machinery activity, and insufficient immune stimulation all lead to low RNAi efficiency

(Reviewed in Cooper et al., 2019). In addition, the secondary structure of a target gene can also render certain regions of the gene unavailable and hence hamper effective

RNAi-based gene silencing (Shao et al., 2007). RNAi results in transient gene silencing, hence, depending on the cell type and concentration of siRNA, duration, time point, the level of gene knockdown could be different (Pancoska et al., 2004).

Figure 1.21. Schematic steps in RNAi (Adapted from Cooper et al., 2019)

Nevertheless, RNAi is still a useful tool in both fundamental and applied research.

With the availability of recent genome editing tools such as CRISPR/Cas (clustered regularly interspaced short palindromic repeats/Cas), some of the weakenesses of the

RNAi approach can be circumvented. For example, rather than transient mRNA knockdown to study gene functions, they could be completely knocked out through genome editing (Cooper et al., 2019; Le Trionnaire et al., 2019). A genome editing protocol in aphids was recently reported (Le Trionnaire et al., 2019; Tyagi et al., 2020).

This is a valuable tool that will greatly facilitate studies looking at interactions between aphids and viruses, and beyond.

1.4.8. Statistical methods and calculation bottlenecks

Statistical methods together with biological assays have been used to correctly

interpret the appropriate results of scientific natural investigations. Using genetic variations and a narrow pathway to discriminate between different individual is a biological method that is used to estimate and quantify the populations. Genetic bottlenecks can occur in virus populations when only a few variants transmit horizontally from infected host to non-infected plants. Moury et al. (2007) used virulent and avirulent PVY populations on pepper plants and transmission by M. persicae as bottleneck phenomenon to quantify the initial transmitted virus particles by a single aphid. They found that most single aphids transmitted 0.5 to 3.2 PVY particles to the plants. In another NP virus (CMV) and using different genotype of CMV belonging to subgroups IA and II and aphid transmission bottleneck, the quantity of transmitted

CMV by individual aphid as founder virus in un-infected plants is 1 to 2 particles

(Betancourt et al., 2008). Furthermore, Betancourt and team believed that a horizontal transmission bottleneck could cause a random genetic drift in the CMV population, resulting losing some genes in the aphid population.

Chapter 2

Potato virus Y (PVY) strains in Belgian seed potatoes and first molecular detection of the N-Wi strain

This chapter was published in:

Bahrami Kamangar S, Smagghe G, Maes M, De Jonghe K (2014) Potato virus Y

(PVY) strains in Belgian seed potatoes and first molecular detection of the N-WI strain. Journal of Plant Diseases and Protection 121: 10-

19. https://doi.org/10.1007/bf03356485

Chapter 2

Potato virus Y (PVY) strains in Belgian seed potatoes and first molecular detection of the N-Wi strain

2.1. Introduction

Potato virus Y (PVY) is a major pathogen in potato production worldwide. The two main strain groups O and N are responsible for most of the damage in potato growing areas (Kerlan, 2006; Rolot, 2009). Apart from O and N, PVYC, PVYZ and PVYE are also recognized as PVY strain groups (Karasev and Gray, 2013). Obvious mosaic, stunting and leaf drop are reported as the main symptoms of the O strain, whereas the N strain causes mild mosaic and occasional leaf necrosis (Singh et al., 2008). Over the past years, a lot of progress has been made in understanding the molecular properties of PVY strains. PVYO and PVYN are considered to be the two starting groups from which one to three recombinations resulted in several subgroups (Nie et al., 2013). Within the N strain group, the NTN strain (PVYNTN) is the causal agent of the potato tuber necrotic ringspot disease (PTNRD) (Beczner et al., 1984; Le

Romancer et al., 1994). In some potato cultivars, this strain can cause severe damage, both on leaves and tubers, whereas recombinants belonging to another N-like subgroup containing the Wilga strain (PVYN-Wi) (Chrzanowska, 1991; Glais et al.,

2001) and PVYN:O (Singh et al., 2003) mainly produce mild symptoms and can even remain latent in potato. However, symptoms caused by the same PVY strains can vary a lot depending on the potato cultivar (Nie et al., 2012). Molecular studies have revealed that even within the subgroups considerable variations in the PVY genome evolved differently, e.g. in North America and Europe, resulting in a very complex

situation of strains, strain groups and variants (Karasev and Gray, 2013). However, these variations are mainly introduced by recombination in different parts of the genome

(Glais et al., 2002; Nie and Singh, 2003a; Lorenzen et al., 2006a; Chikh Ali et al.,

2007a, b; Chikh Ali et al., 2010b). Three to four recombinant junctions in NTN strains and only one or two recombinant junctions in PVYN-Wi and PVYN:O have been reported

(Glais et al., 2002; Nie and Singh, 2003a; Lorenzen et al., 2006a, b; Schubert et al.,

2007; Visser et al., 2012; Karasev and Gray, 2013). Position and length of these recombinant segments, resulting in variations in recombinant junctions or breakpoints, have led to several publications on the existence of NTN variants (a and b) and on differentiating N-Wi and N:O subgroups, which have been summarized in the recent reviews of Visser et al. (2012) and Karasev and Gray (2013). Yet, fitness pressure selects and boosts the relative importance of some populations that have been created by point mutations and/or recombination. Geographic isolation could conduct different fitness pressure and natural selection to select and boost various recombinations, resulting in separate lineages, such as some O and Wilga strains in North America (the so called PVYO-O5 subgroup and PVYN:O) (Singh et al., 2008) and NA-PVYNTN that has evolved from NA-PVYN by mutation rather than recombination (Nie and Singh, 2003a).

Available data indicate that the N strain group (including PVYN, PVYNTN and PVYN-Wi strains) is the most common strain group in Europe (Weidemann, 1988; Glais et al.,

1998; Glais et al., 2002; Nie and Singh, 2003b), while in North America, the ordinary strain group (PVYO) still has a predominant place (Crosslin et al., 2006, Lorenzen et al., 2006a, Nanayakkara et al., 2012a). Both in Germany (Lindner, 2008) and

Switzerland (Rigotti et al., 2011), the NTN-strain has been shown to be the most prevalent PVY strain, although both authors report a gradual shift over the years toward prevalence of the PVYN-Wi strain. This shift towards the Wilga strain was also

reported in the Netherlands (Van der Vlugt et al., 2008, Verbeek, 2009) and France

(Kerlan et al., 1999). In France, the presence of Wilga, N, C, Z and O strains were reported, with O as the most dominant strain (Kerlan et al.,1999), but the spread of

NTN in France was also reported recently (Blanchard et al., 2008).

The importance of the N strain for Belgium has also been demonstrated by Rolot and

Steyer (2008). In a more recent study, Rolot (2009) confirmed an increase in importance of the N strain group. Note that all of these records are based on serological tests that cannot differentiate between N and NTN. Additionally, PVYN-Wi is serologically identified as an O strain (Singh et al., 2008). PVY clearly has a large economic impact on potato seed production, but the various strains have a different impact depending on the symptoms they cause. Obtaining accurate data on the occurrence and prevalence of the different strains is thus important. The overall objective of this study is to support the seed potato certification scheme by means of a detailed characterization of the variability in PVY strains in Belgium and their relative importance.

2.2. Materials and Methods

2.2.1. PVY screening and collection of PVY isolates

The survey was initiated with seed potatoes sampled on farms during the Belgian official certification scheme in 2010. 54 potato lot samples (54 farms) were collected from different geographical locations in Belgium (Table 2.2). PVY incidence was determined using the grow-out process in greenhouse, followed by the DAS-ELISA (Clark and

Adams, 1977) test by polyclonal antibody (DSMZ, Germany) on 2700 (50x54) leaf samples collected from individual plants of the tubers. These plants were also were assessed for symptom development. Individual PVY-infected plant samples were stored at -70°C for further analysis.

2.2.2. Strain determination

Total plant RNA was extracted from maximum 100 mg potato fresh leaf material using the Spectrum Plant Total RNA extraction Kit (Sigma, Bornem, Belgium). RNA yield and quality were checked spectrophotometrically (ND-1000 Spectrophotometer,

NanoDrop, Isogen, Temse, Belgium).

First strand DNA was synthesized by adding 1 μl of RNA template to 19 μl of a mixture containing 1 μl of RevertAid Premium Reverse Transcriptase, 4 μl of 5x RT buffer, 1 μl of dNTP Mix (10 mM), 0.5 μl of Ribolock RNase inhibitor and Oligo-dT primer at a final concentration of 2.5 μM (RT, Thermo Fisher Scientific, Waltham, MA, USA). The RT reaction was done at 42°C for 60 min followed by inactivation of the RT by heating at

95°C for 5 min.

Multiplex PCR for strain differentiation was carried out according to Chikh Ali et al.

(2010a) (primers listed in Table 2.1) with some modifications. PCR was conducted using OneTaq™ DNA Polymerase (Bioke, Leiden, the Netherlands) according to the following program: denaturation at 94°C for 4 min, 30 cycles of denaturation at 94°C for 30 s, annealing at 64°C, 62°C and 60°C in the first, second and third 10 cycles for

30 s, respectively, and extension at 68°C for 5 min.

Table 2.1. Primers used in this study Primer Polarity Sequence (5’–3’) Location Reference Application name n156 + GGGCAAACTCTCGTAAATTGCAG 160–179 Chikh Ali et al. (2010a) Multiplex RT-PCR characterization o514 + GATCCTCCATCAAAGTCTGAGC 515–536 Chikh Ali et al. (2010a) Multiplex RT-PCR characterization n787 – GTCCACTCTCTTTCGTAAACCTC 770–792 Chikh Ali et al. (2010a) Multiplex RT-PCR characterization n2258 + GTCGATCACGAAACGCAGACAT 2260–2281 Lorenzen et al. (2006a) Multiplex RT-PCR characterization o2172 + CAACTATGATGGATTTGGCGACC 2169–2191 Lorenzen et al. (2006a) Multiplex RT-PCR characterization n2650c – TGATCCACAACTTCACCGCTAACT 2627–2650 Lorenzen et al. (2006a) Multiplex RT-PCR characterization o2700 – CGTAGGGCTAAAGCTGATAGTAG 2678–2700 Chikh Ali et al. (2010a) Multiplex RT-PCR characterization S5585m + GGATCTCAAGTTGAAGGGGAC 5578–5598 Lorenzen et al. (2006a) Multiplex RT-PCR characterization o6400 – GTAACTCCTAAACAAATGGTGGTTCG 6405–6430 Chikh Ali et al. (2010a) Multiplex RT-PCR characterization n7577 + ACTGCTGCACCTTTAGATACTCTA 7582–7605 Chikh Ali et al. (2010a) Multiplex RT-PCR characterization YO3-8648 – CTTTTCCTTTGTTCGGGTTTGAC 8635–8657 Schubert et al. (2007) Multiplex RT-PCR characterization SeroN – GTTTCTCCTATGTCGTATGCAAGTT 8864–8888 Chikh Ali et al.(2010a) Multiplex RT-PCR characterization + GGATCCAATTAAAACAACTCAATA 5' end Nie & Singh 2003a Sequencing 1 – CATTTGTGCCCAATTGCC 1091–1073 Nie & Singh 2003a Sequencing 2 + TTCAGTTCTCAAGCGCTGAA 1033–1052 Nie & Singh 2003a Sequencing

– TCTTAGTGAAATCCTTTGCAT 2133–2113 Nie & Singh 2003a Sequencing + GCGATGTTGATTAACATT 2085–2102 Nie & Singh 2003a Sequencing 3 – TTATTGCCTGACACACTGC 3294–3276 Nie & Singh 2003a Sequencing + TCTTCAGGCGTTTGCCAACTTT 3203–3224 Nie & Singh 2003a Sequencing 4 – TTGCGCATCAACAAATGATTGG 4367–4346 Nie & Singh 2003a Sequencing + TGTCAGCTACTCCAGTGGGAAG 4266–4285 Nie & Singh 2003a Sequencing 5 – TCAGTGCGCAATGTGTATGC 5268–5250 Nie & Singh 2003a Sequencing + CACGAAATGCTTTGGGAAAC 5169–5185 Nie & Singh 2003a Sequencing 6 – CAGGAAACTTGGCAATGCCATT 6198–6177 Nie & Singh 2003a Sequencing + GCCACACAACCCACTCAAA 6143–6161 Nie & Singh 2003a Sequencing 7 – TCCGACTGCAGCTTTCAT 7400–7383 Nie & Singh 2003a Sequencing + GGCTATCAATAGGGTTATCAT 7289–7309 Nie & Singh 2003a Sequencing 8 – CTTGATGGTGCACTTCATAAG 8571–8551 Nie & Singh 2003a Sequencing + GCTTTCACTGAAATGATGGT 8502–8521 Nie & Singh 2003a Sequencing 9 – GTTTTCCCAGTCACGACTTTTTTTTTTT 3´ end Nie & Singh 2003a Sequencing 5Pr – ATCTGGACATCAGTCTTGTATC This study Sequencing + TGTCAACCAAAGGAGGGTCTG This study Sequencing Gap 1 – ACCGGTTCAGTTAAGTGCTCT + AAATTGATCCAGCGAAGGGC This study Sequencing Gap 2 – CTCCTCCTTCTCTGAAAGGTGA + ATGGTGTTGCAAGTTGTTAAGAA This study Sequencing Gap3 – CAGCATCGAACACCATGATGA + TTCAAGAAGCCAACACTGCG This study Sequencing Gap 4 – GCAATGCTCCTATTGTCAATGTC + AACTTGCCAGTGATGACAGG This study Sequencing Gap 5 – GCGCCTGCTATGATCAAGTC + GGTATGGGCAAGTCAAGCAG This study Sequencing Gap 6 – ACAGGGAAATCTTTCGGCAT + ACAGGGAATTTGCAAGCTGT This study Sequencing Gap 7 – TCTAAAGGTGCAGCAGTGAA + GACAGCACGTGTGTATTCTTTG This study Sequencing Gap 8 – CTGTGATTGAGTTGCTCGAGT 3Pr + GACAGCACGTGTGTATTCTTTG This study Sequencing

This was followed by a final 5 min extension step at 72°C. PCR products were visualized using Qiaxel capillary electrophoresis (QIAxel Advanced System). The obtained amplicons were compared to the expected size (Chikh Ali et al., 2010a) and those of the

PVY reference isolates that were included in the experiment. The whole genome of the nine isolates was compiled (assembled using Codoncode and MEGA5 software) after amplifying nine segments and gaps using the primer pairs described by Nie and

Singh (2003a) and also designed in this study. Extra primer pairs were developed for additional sequencing of the overlapping regions of the nine segments (Table 2.1).

PCR product purification and sequencing with the corresponding forward and reverse primers were performed by Macrogen Inc. (Amsterdam, the Netherlands). Consensus

sequences of sense and antisense strands of all nine selected isolates were aligned and were deposited in GenBank and the accession numbers were linked to the isolate overview in Table 2.2. A phylogenetic dendrogram was compiled by means of

Neighbour-joining test, test of phylogeny (with 1,000 bootstrap replicates). The selected

PVY isolates of this study were compared with a representative number of PVY isolates of the respective strain (sub) groups that were retrieved from the GenBank database (Table 2.3). A schematic diagram of the recombinant junctions (breakpoints) was drafted based on the RDP4 multiple recombination detection software (Martin et al., 2010).

2.3. Results

2.3.1. Prevalence of PVY types in Belgium

An overall general PVY infection rate of 3.6%, ranging from 0% to a maximum of 42%, was recorded in the 54 seed potato lots that were included in the survey. PVY infected tubers were identified in 42.5% (23/54) of the seed lots. A geographical distribution of samples positive for PVY, covering 23 administrative divisions in Belgium, is presented in Table 2.2.

Table 2.2. Sampling location, infection rate and PVY strain type for different potato cultivars to PVY in Belgian potato seed farms based on ELISA screening and RT-PCR strain typing. For nine selected isolates, the isolate ID and GenBank accession numbers are added

Isolate ID and No. of farms % Infected seed Location Cultivar Strain (RT-PCR) GenBank tested (total 54) lots (ELISA) Accession No. NTN Adinkerke 1 16 Eersteling Mix (NTN&O&N-Wi) 6 Rode eersteling Unknown Amberloup 2 4 Spunta NTN 0 Draga --- Assenede 3 0 Draga --- 0 Draga ---

0 Marfona --- Boekhoute 3 0 Marfona --- 8 Paramount NTN x2 Brielen 1 0 Granola --- Ehein-Neupré 1 2 Spunta NTN Gistel 1 0 Spunta --- Grimbergen 1 0 ------0 Diamant --- Hoeleden 3 0 Diamant --- 0 Diamant --- 4 Kennebec NTN Hompré 2 10 Spunta NTN x2 JQ969033 Kieldrecht 1 0 Granola --- Koekelare 1 0 Desiree --- Langemark- 4 Spunta NTN 2 Poelkapelle 0 Marfona --- 0 Diamant --- Lo-Reninge 3 0 Cara --- 2 Cara NTN Méeffe 1 0 Diamant --- 8 Diamant N JQ969036 2 Diamant NTN JQ969037 Milmort 4 2 Spunta NTN 4 Spunta NTN 0 Spunta --- 0 Spunta --- Moerbeke- 0 Draga --- 6 Waas 0 Draga --- 0 Draga --- 0 Draga --- 0 Marfona --- 0 Marfona --- 0 Spunta --- 10 Spunta NTN x2 Roeselare 8 JQ969035 14 Spunta NTN x2 18 Anosta NTN x2 & N-Wi* JQ969040* 10 Anosta N-Wi JQ969039 2 Spunta NTN x2 Sommière 1 0 Spunta ---

14 Lady Rosetta O x3 JQ969038 Tinlot 2 0 Hermes --- 0 Nicola --- 0 Diamant --- Veurne 5 0 Charlotte --- 4 Nicola N-Wi JQ969041 2 Charlotte Unknown Vlissegem 1 6 Spunta NTN x2 Wulpen 1 42 Bintje NTN x7 JQ969034

2.3.2. Strain determination

Forty PVY isolates were selected from PVY infected samples in ELISA tested from 21 locations in Belgium. They were subjected to strain determination using the multiplex RT-

PCR protocol described by Chikh Ali et al. (2010a). As shown in Figures 2.1 and 2.2, analysis of the obtained fragments confirmed the presence of four strains and a variation (a and b) in NTN strains. These isolates were identified as NTN, O, N-Wi, and N with a relative abundance of 75%, 7.5%, 7.5%, and 2.5%, respectively. A minor percentage (2.5%) of the infected plants revealed mixed infections with two or three strains (NTN and N-Wi and/or O), and 5% of the isolates could not be identified, probably also due to mixed infections (Fig. 2.1).

Table 2.3. Sequences retrieved from GenBank and used in phylogenetic analysis

GenBank Accession No. Strains Country

JQ969033, JQ969035, JQ969037, JQ969034, JQ969039, JQ969040, JQ969041, NTN, N-Wi, N, O Belgium JQ969036, JQ969038

JQ924285, JF928458, JF928459, JF928460 O, E, NTN Brazil

AY166867, AY166866, U09509 O, N, NTN Canada

HM590407 O China

HM991454, AJ890348 C, N-Wi France

HE608964, AJ889868, HE608963, AJ890346, NTN, N-Wi Germany AJ890347, AJ890350

JF927749, JF927761, JF927756, JF927752 NTN Hungary

AB711154, AB711150, AB711149, O, N Japan AB714135 AM268435 N New Zealand

EU563512 C the Netherlands

JF795485, AJ890343, AJ890342, AJ889866 NTN, N-Wi, N Poland

X97895 N Switzerland AB270705 N Syria AJ585195, AJ585198, AJ585197, JX424837, HQ912892, HQ912891, HQ912897, HQ912914, HQ912913, HQ912884, O, N United Kingdom HQ912883, HQ912909, HQ912879, HQ912876, HQ912878, HQ912875 HQ912874, HQ912868, HQ912896, HQ912863, HQ912871, HQ912862, HQ912872, HQ912870, HQ912869, O, O5, N-Wi, N:O, NTN, NA,PVY- United States of HQ912890, HQ912893, HQ912864, N America FJ643478, EF026074, EF026076,

FJ204166, FJ204165, FJ204164, EF026075, EF026074, AY884983, AY884984 NC_004039 Potato virus A Hungary

NC_004010 Potato virus V United Kingdom

M96425, NC_001517 Pepper Mottle virus United States of America

The obtained virtual banding patterns of the PCR products of a selection of representative PVY isolates and reference isolates (JKI, Germany) produced with the six primer pairs of the multiplex RT-PCR is presented in Fig. 2.2. Phylogenetic analysis of the sequences (Fig. 2.3) was in a good correspondence with the multiplex banding pattern, except for GBVC_PVY_15 NTN (JQ969034).Based on the banding patterns in the multiplex PCR, three isolates in this study were identified as Wilga strains

(GBVC_PVY_26 N-Wi (JQ969039), GBVC_PVY_23 N-Wi (JQ969040) and

GBVC_PVY_34 N-Wi (JQ969041)).

hausen 1”.

Fig. 2.3. Phylogenetic analysis of PVY isolates from Belgian seed potato production (this study) (▲). All reference sequences are retrieved from GenBank (http://www.ncbi. nlm.nih.gov). Statistical Methods: Neighbour-joining test, test of phylogeny (with 1,000 bootstrap replicates), evolutionary analyses were conducted with MEGA5. The bars represent schematic recombination maps of the respective strain groups.

The phylogenetic analysis confirmed the attribution to the Wilga subgroup and revealed that all three isolates cluster in the N-Wi subgroup (Nie and Singh 2003b, Karasev et al., 2011, Karasev and Gray 2013, Visser et al., 2012). However, the variability between the Wilga strain (GBVC_PVY_23 N-Wi (JQ969040) and GBVC_PVY_26 N-Wi

(JQ969039)) from two infected Anosta lots from the same location (Roeselare) was also significant with both isolates clustering in different clades (Fig. 2.3). Additionally, the recombination pattern of all three Wilga strains was similar showing two breakpoints at

RJ1 and RJ2 which is corresponding to the other N-Wi strains of which the complete sequence is available in GenBank (Fig. 2.4).

Four isolates that were identified as NTN strains by means of the multiplex PCR were also fully characterized. The multiplex PCR banding pattern (Fig. 2.2) produced two bands (441 and 1307 bp) for GBVC_PVY_15 NTN (JQ969034) and three for other

NTN isolates. Based on this difference, Chikh Ali et al. (2010a) differentiated the isolates into type A (GBVC_PVY_3 NTN (JQ969035), GBVC_PVY_9 NTN

(JQ969037) and GBVC_PVY_37 N_NTN (JQ969033)) and type B (GBVC_PVY_15 NTN

(JQ969034)). However, based on the full genome sequence analysis (Fig. 2.3), the

Belgian isolates were all identified as NTNa (type A) as described in the review of

Karasev and Gray (2013), clustering in a major clade with other NTNa strains. The

NTNb reference isolates which were retrieved from GenBank cluster in a separate small clade, also contained the NTN strain AJ889866 (Schubert et al., 2007) from

Poland. Additionally, this was also confirmed by the recombination map (Fig. 2.4) of the NTN isolates in this study that showed three breakpoints, corresponding to RJ2,

RJ3 and RJ4, and identified all of them as NTN type A after Karasev and Gray (2013).

The alleged O strain (GBVC_PVY_20 O (JQ969038)) also clustered in the same group together with O strains from US, EU and Japan that were deposited in GenBank (Fig.

2.3). Additionally, the suspected N (GBVC_PVY_10 N (JQ969036)) clustered with the corresponding N strains that are available in GenBank, including N Mont (AY884983), N

605 (X97895) and N New Zealand (Fig. 2.3). PVYE, PVYC strains and other potyviruses

(PotAto virus A and V and Pepper Mottle virus) were also included in the phylogenetic analysis and clustered significantly different from all isolates (Fig. 2.3).

Fig. 2.4. Schematic recombination map of nine selected PVY isolates compiled by the RDP 4 multiple recombination detection software. RJ: recombination junction.

2.3.3. Symptoms, strain and potato cultivar relation

The seed potato samples infected with the Wilga strain belonged to the cultivars

Anosta and Nicola were collected in Veurne and Roeselare (Table 2.2). The most

prevalent strain (NTN) was detected in the following cultivars: Spunta, Diamant, Bintje,

Eersteling, Anosta, Paramount, Cara and Kennebec.

PVY-infected plants ranged from symptomless to mottling, mild and severe mosaic and leaf malformation. Tissue necrosis and stunting were only occasionally observed.

Cultivar ‘Spunta’, produced on 21 farms, was found to be mainly infected with NTN and showed symptoms ranging from mild mosaic to leaf necrosis. N mainly caused mosaic, while O and N-Wi symptom development was more variable, ranging from symptomless to mottling, mild to severe mosaic and growth reduction on several cultivars. No clear correlation between the symptoms and the cultivars could be recorded.

2.4. Discussion

Based on the ELISA screening results, the average of PVY infection rate of 3.6% in

Belgian seed potato farms in our study was slightly higher than the rate (2.4%) obtained in 2007 (Rolot, 2009). In agreement with the report of Rolot (2009), our results also confirmed that the N strain group (NTN, N-Wi and N strains) is the most prevalent (89.5%) in Belgium. In accordance with published data from other important

West European potato growing areas, this study confirmed the relative importance of the NTN strain compared to the specific N strain. However, the percentage NTN strains that was detected (75%) was lower than what was recorded in Germany (Lindner,

2008), the Netherlands (Verbeek, 2009) and Switzerland (Rigotti et al., 2011).

The NTN population collected in Belgium did not consist of homogenous isolates based on analysis using the PCR method of Chikh Ali et al. (2010a). Presence or absence of the 633 bp amplicon is discriminative between NTN variants (A and B) and indicates the presence or absence of an extra recombinant junction (RJ1) at the 5´end. NTNb therefore starts with a small segment of the O strain (Chikh Ali et al., 2010a, Karasev

and Gray, 2013). GBVC_PVY_15 NTN (JQ969034) lacked this 633 bp amplicon, but sequence analysis revealed that this was rather due to a base replacement at a critical position (Fig. 2.5) at the annealing site with the relevant forward primer (n156) (Chikh

Ali et al., 2010a), rather than a recombination on which the actual A and B typing is based (Karasev and Gray 2013). We recommend that future sequence analysis on sufficiently large fragments of the genome in the RJ regions allowing to reconstitute recombination maps, or ideally, full genome sequencing is necessary to accurately type PVY isolates.

Fig. 2.5. Comparison of sequence of annealing sites of the NTN strains with relevant forward primer (n156) (Chikh Ali et al., 2010 a). Base variations are indicated in black squares.

Our results confirmed the presence of N-Wi strain in Belgian seed potatoes for the first time. In addition, the phylogenetic tree also showed that the Belgian Wilga isolates clustered in the different interior branch of the European and American Wilga isolates

(Fig. 2.3) (Glais et al., 2002, Nie and Singh, 2003b). Additional pathological and biological characterization of these isolates is necessary to further type the NTN and N-

Wi populations. This would also allow researchers to further categorize the NTN strain into PVYNTN- Hun (European PVYNTN) and PVYNTN-Tu 660 (North American

PVYNTN), the two NTN types identified and discussed by several research groups

(Thole et al., 1993, Nie and Singh, 2002, 2003a, b, Piche et al., 2004, Chikh Ali et al.,

2007b, 2010b, Hu et al., 2009, 2011).

In summary, this study presented the relative distribution of the PVY strains across seed potato lots in Belgium. We could conclude that NTN is the most dominant strain in all

parts of Belgium and that the NTN isolates belong to the NTNa type. Additionally, the presence of the Wilga strain was confirmed and genetically fully characterized for the first time in Belgium. All Wilga strains belong to the N-Wi subgroup and no N:O variants were identified. The other strains, O, and PVY-N, appeared less important and were also more restricted in their geographical distribution. No PVYC, PVYE or PVYZ was put in evidence.

Chapter 3

Quantity and transmission efficacy of an isolate of the Potato virus

Y-Wilga (PVY N-Wi) by aphid species reared on different host plants

This chapter was published in:

Bahrami Kamangar S, Taning CN, De Jonghe K, Smagghe G (2019) Quantity and transmission efficiency of an isolate of the potato virus Y–wilga (PVYN−Wi) by aphid species reared on different host plants. Journal of Plant Diseases and Protection, 126:

529-534. https://doi.org/10.1007/s41348-019-00266-0

Chapter 3

Quantity and transmission efficacy of an isolate of the Potato virus

Y-Wilga (PVY N-Wi) by aphid species reared on different host plants

3.1. Introduction

Potato virus Y (PVY), the type species of potyviruses, is a well-known destructive and worldwide occurring virus and infects a group of different crops, particularly potato. PVY is transmitted in a non-persistent manner by more than 50 different specific aphid vector species (de Bokx and Huttinga, 1981). Information on the factors that influence transmission efficiency is important in epidemiology and virus management. Different factors that affect virus transmission include the aphid species, virus ‘species’ or strain

(Pelletier et al., 2008; Verbeek et al., 2010), aphid feeding behavior (Nanayakkara et al., 2012b), both source and recipient host plant of the virus (Cervantesa and Alvarez,

2011), environmental factors (temperature and humidity) (Singh et al., 1988) and aphid behavior (Powell et al.,1995). Among all factors which could possibly explain the reported variability in PVY transmission, the effect of the plant host species on which the aphid develops, on the ability of the aphid to acquire and transmit the virus, has received the least attention.

Another important parameter which warrants more investigation is the quantification method for PVY, for identifying efficient aphid vectors. So far, the direct molecular quantification of potyviruses has only been conducted in whole bodies of single aphids

(Singh et al., 1996; Cambra et al., 2004; Olmos et al., 2005; Moreno et al., 2007) and not in stylets for which reports are limited to detection (Pelletier et al., 2012; Boquel et al., 2013; Zhang et al., 2013). Based on information provided by the Electrical

Penetration Graphs (EPG) technique (Martin et al., 1997; Powell 2005), it is known that

the inoculation of non-persistently transmitted viruses by aphids is associated with the active injection of saliva into the cytoplasm from the common duct during the sub- phase (II-1). The common duct covers about 1% of the stylet length (Forbes, 1969), and based on electron microscopy investigations, there are more viruses retained in the food canal of aphid vectors (Ammar et al., 1994). In addition, GFP-labeled helper component microscopy showed dense fluorescence illumination in the common duct or acrostylet in the distal part of aphid stylets (Uzest et al., 2010). This distal stylet part could be an efficient and important location in aphids for the retention and may be also transmission of some semi- or non-persistently transmissible plant viruses, such as Caulimovirus and Potyvirus species, respectively (Fereres 2007). In this study, the main objectives were designed to investigate the effect of different host plants on which aphids were reared on their ability to acquire and transmit PVY, and to determine whether the quantity of PVY in the aphid vector stylet could be used as a useful trait for the evaluation of vector efficiency. As such, the transmission efficiency and quantity of a PVYN−Wi isolate was measured in clones of both vector and non- vector aphids (stylet alone or intact whole bodies with stylet). Then, the existence of a correlation between the quantity of the virus in the stylet of the aphid or their intact whole body and the virus transmission efficiency was evaluated. The findings could be useful in epidemiological studies or predictive schemes.

3.2. Material and Methods

3.2.1. Aphids, plants and virus isolate

M. persicae clones (A and B) and Aphis fabae Scopoli (1 clone) were collected from zucchini fields, and Aulacorthum solani (1 clone) from potato plants, both from around

Ghent, Belgium. Acyrthosiphon pisum was obtained from an existing colony culture at

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the Laboratory of Agrozoology, Ghent University. All aphid colonies were maintained as clones. For the aphid species transmission efficiency bioassays, all aphid colonies of M. persicae, A. fabae, A. solani and A. pisum were started from individuals and maintained on virus symptom-free broad bean plants (Vicia faba L.). In addition,

M. persicae clones (B) were reared on Capsicum annum L. (pepper), Brassica rapa L.

(Brussels sprout), V. faba (broad bean) and Solanum tuberosum L. (potato) to determine the effect of host (rearing) plant on the transmission efficiency. In all of the experiments, a Potato virus Y–Wilga isolate (GBVC_PVY_26; GenBank Accession

N−Wi No. JQ969039 PVY ), originally collected from a potato farm in Belgium (Bahrami

Kamangar et al., 2014), was used. This strain was maintained on 30–60 day old potato plants (Anosta cultivar) throughout the study, and the youngest leaflets were used as the virus source in the virus transmission bioassays. The concentration of the virus in these leaves was determined by RT-qPCR. Nicotiana benthamiana seedlings used as indicator plants for aphid transmission tests were grown in a growth chamber (25 °C and 16:8 h Light: Dark) and inoculated with the PVYN−Wi isolate at second leaf stage by different aphids.

3.2.2. RNA‑transcript synthesis

A 150 bp-PVY nucleotide sequence on the target region (NIb) was amplified using the PVY 100–5 FP/100–5 RP primer pair (Agindotan et al., 2007). The fragment was inserted into the pJET1.2/blunt cloning vector (Thermo Fisher Scientific, Waltham,

CA) and cloned into TOP10 Escherichia coli electrocompetent cells. Transformed bacteria were cultured on Super Optimal Broth medium with Catabolite repression

(SOC) and a colony qPCR test with the same primer set confirmed transformation.

Extracted plasmids using the QIAprep Spin Miniprep Kit (Matrix Technologies, Kansas

City, KS) were linearized by the Fast Digest restriction enzyme XbaI, and

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subsequently, transcripts were synthesized by means of the MEGA script T7

Transcription Kit (Thermo Fisher Scientific, Lafayette, CO). Template DNA was removed by a TURBO DNase step. The concentration (ng/µL) of the purified RNA transcripts (transcript size: 150 viral and 56 vector nucleotide) was measured using

NanoDrop ND-1000 spectrophotometer (Isogen Life Science, De Meern, the

23 Netherlands), and the Avogadro constant (6.023 x 10 molecules/mol) was used to

–10 eventually estimate the virus copy number. A tenfold dilution series up to 10 was prepared for use as a standard [slope = − 4.216, y-intercept = 46.55, efficiency (R2) =

0.993] in the quantification tests.

3.2.3. PVYN−Wi transmission bioassays

Two transmission bioassays were set up. In the first setup, the transmission efficiency of PVYN−Wi by both efficient and non-efficient aphid vectors which were reared on the same host species (broad bean) was assessed. Twelve apterous adults of either

M. persicae (clone A), A. fabae, A. solani or A. pisum in three replications (12 aphids x 3 replications) were aspirated and starved for 3 h to prompt probing behavior in an acquisition feeding step. The starved aphids were given an acquisition access period

(AAP) of 5 min on PVYN−Wi-infected detached potato leaves and were then individually transferred to single N. benthamiana seedlings, which were placed in a small insect proof cage. After 24 h, the aphids were killed using pirimicarb (Pirimor).

Three to four weeks later, symptom development was assessed and evaluated by means of DAS-ELISA (Clark and Adams 1977). The data were subjected to analysis of variance (ANOVA) and Duncan test with the SPSS software (IBM, Armonk, NY). The aphid species, A. fabae, A. solani and A. pisum, were used in this bioassay because they have been previously described as non to less efficient aphid vectors, while M.

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persicae (clone A) represents a well-described efficient aphid vector (Van Hoof, 1980;

Piron, 1986; Verbeek et al., 2010; Boquel et al., 2011a; Mello et al., 2011).

In a second transmission bioassay, the effect of C. annum (sweet bell pepper), B. rapa

(Brussels sprouts), V. faba (broad bean) and S. tuberosum (potato) as different host plants, was assessed for PVYN−Wi transmission efficiency by M. persicae (clone B).

M. persicae was maintained on each of the host plants starting from a single aphid.

Of each colony maintained on the four types of host plants, 12 aphids were subjected to a transmission experiment. In a similar setup as in the first transmission bioassay,

M. persicae aphids from each host plant were starved, given an acquisition access period on PVYN−Wi-infected detached potato leaves and then transferred to N. benthamiana seedlings. After 24 h, the aphids were killed and PVYN−Wi transmission was assessed as described in the first transmission bioassay.

3.2.4. PVY N-Wi quantification in the aphid stylet and intact whole body

Based on the results obtained from the transmission bioassays, the stylet and the intact whole body of a non-efficient aphid vector species from the first transmission bioassay (as control) and from the least and most efficient M. persicae vector as kept on the different tested host plants in the second transmission bioassay were subjected to PVYN−Wi quantification tests. A one-step probe-based real-time RT-qPCR, using the AgPath-ID One-Step RT-qPCR kit (Thermo Fisher Scientific), was applied. The thermal cycling condition was set at 45 °C for 10 min, 95 °C for 10 min and 40 cycles of 95 °C for 15 s, 50.2 °C for 30 s and 60 °C for 45 s. The absolute PVYN−Wi copy numbers in individually dissected stylets and intact whole bodies (12 for each treatment) were estimated using a tenfold serial dilution of the standard transcripts in the one-step RT-qPCR.

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3.2.5. Preparation of aphid and plant samples, and RNA extraction

Single stylets or complete bodies of aphids were subjected to RT-qPCR to evaluate the quantity of PVY. Aphids were starved (3 h), and after a 5-min AAP on detached infected potato leaves, they were anesthetized by chilling for 5 min at -4 °C. Four stylets from four aphids were dissected individually in each three replications (3 x 4 individual), using micro-scissors (5 mm blades, 0.1 mm Tips, PN: 14,003, World

Precision Instrument, Sarasota, FL) under a stereomicroscope (Leica S8 APO,

Wetzlar, Germany). The stylets were cut from the basal segment of the labrum.

Dissected stylets (12 individuals) or intact whole bodies (12 individuals) of aphids were individually placed into 1.5-mL tubes and immediately immersed into liquid nitrogen. The above-mentioned samples were crushed and lysed in RLT buffer

(RNeasy Plant Mini kit, Qiagen, Venlo, the Netherlands). The samples were then

N-Wi subjected to the RNA extraction protocol of the RNeasy kit. RNA of PVY - infected and healthy potato leaves (100 mg) was extracted using the same kit and was included as control in each RT-qPCR experiment.

3.3. Results

3.3.1. RNA transcript and standard

The transcript size of PVY is 206 nucleotides (150 + 56 viral and vector). The copy number was calculated based on the molecular weight (66191.4 g/mol), corresponding

8 2 −3 −9 to a range from 3.1E to 3.1E transcript copies or 1.3E to 1.3E ng/µL (PVY) for the serial dilution that was included in the RT-qPCR assays of this study (Table 3.1).

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3.3.2. Transmission of PVYN−Wi by aphids

Among the four aphid species, only M. persicae (clone A) transmitted PVYN−Wi with an efficiency of 39 ± 3%, while A. pisum, A. fabae and A. solani clones did not transmit

PVYN−Wi (Fig. 3.1). A comparison of the means of PVYN−Wi transmission efficiency by

M. persicae (clone B) reared on different plants demonstrated that the transmission was significantly higher (81 ± 10%) when the aphid host was Brussels sprouts compared to broad bean (50 ± 8%) and the means were grouped as (b) and

(a), respectively (0.05 < p value < 0.1; Duncan). Sweet pepper (69 ± 17%) and potato

(67 ± 8%) as host plant species resulted in intermediate (ab) transmission efficiencies

(Fig. 3.2). A. solani as non-vector (control) and M. persicae (clone B) reared on

Brussels sprouts and broad bean, as the respective most and least efficient vector, were selected for PVYN−Wi quantification in the aphid stylet and intact whole body.

Table 3.1. Calculated concentrations of a serial dilution of transcripts and ranges of virus quantity in aphid stylet and intact whole body. Each sample contains one dissected stylet or one intact whole body. *The ranges are derived from maximum and minimum of all samples. The detailed data are presented in Fig. 3.3. (The standard curve parameters: slope = − 4.216, Y-intercept = 46.56 and efficiency = 0.993).

Cq-value (means or RNA concentration Virus copies per sample Sample contents ranges) (ng/µL) (means or ranges)

−3 8 Transcript × 10–4 10.43 1.34E 3.1E

−4 7 Transcript × 10–5 15.6 1.34E 3.1E

−5 6 Transcript × 10–6 19.7 1.34E 3.1E

−6 5 Transcript × 10–7 22.4 1.34E 3.1E

−7 4 Transcript × 10–8 26.8 1.34E 3.1E

−8 3 Transcript × 10–9 32.8 1.34E 3.1E

−9 2 Transcript × 10–10 35.9 1.34E 3.1E

−7 4 M. persicae stylet* Not assigned–29.16 Not assigned–2.72E 0–1.33E

−9 −7 3 5 M. persicae intact whole body* 32.94–24.3 7.81E –9.42E 1.71 E –2.234E

−3 9 Leaf tissue of Infected potato 7.8 6.7E 1.54 E

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N−Wi Fig. 3.1. Transmission efficiency of PVY isolate by clones of different apterous viviparous females of aphid species (mean ± SD). (Each treatment included 12 aphids x 3 replications)

Fig. 3.2. Transmission efficiency of PVYN−Wi isolate by M. persicae (clone B) (mean ± SD), reared on different host plants, broad bean (Vicia fabae), potato (Solanum tuberosum), pepper (Capsicum annum), Brussels sprout (Brassica rapa). Means labeled by the same letter are not significantly different with 0.05 < p < 0.1. (Each treatment included 12 aphids x 3 replications)

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3.2.3. Quantification of PVYN−Wi in the aphid stylet and intact whole body

N−Wi The quantification of PVY in A. solani (non-vector) kept on broad bean, M. persicae (clone B) on broad bean (median efficient vector) and M. persicae (clone B) on Brussels sprouts (most efficient vector) was performed by means of a one-step RT-

3 qPCR. In the stylet of M. persicae on broad bean, the amounts of PVY (1.1 ± 2.1E copies) were significantly less (p < 0.01) than in M. persicae kept on Brussels sprouts

3 (4.6 ± 2.6E copies), whereas PVY was not detected in the stylets of A. solani (Fig.

3.3A). Additionally, the quantity of PVY in the intact whole body of A. solani (67.8 ±

3 3 83.1E copies) and M. persicae (62.8 ± 58.7E copies) was nearly equal, while the estimated PVY quantity in whole bodies of M. persicae on Brussels sprouts (13.3E ±

3 8.7E copies) was the lowest (Fig. 3.3B). The percentage of transmission correlated

Fig. 3.3. Amount of PVYN−Wi copies (mean ± SD) detected in the stylet (A) and intact whole body (B) of A. solani clones (non-vector aphid) and M. persicae (clone B) when reared on either broad bean or Brussels sprouts. Means labeled by the same letter are not significantly different with p > 0.01. (4X3 samples for each treatment).

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directly with the amounts of the virus in the stylet of the aphid (r = 0.7; p < 0.05), but there was no correlation with the virus amounts in the intact whole bodies (r = 0.06; p > 0.05).

3.3. Discussion

Understanding the factors that can influence the transmission efficiency of a virus is very important in the management of the virus in question and in epidemiology. An important factor that has received the least attention is the effect of the host plant of the aphid on the transmission efficiency of PVY. In this study, clones of four aphid species (A. fabae, A. solani, A. pisum and M. persicae) were reared on non- infected broad bean, exposed to PVY-infected potato leaves and later introduced to N. benthamiana seedlings, as indicator plants for aphid transmission tests. Out of the four aphid species tested, three (A. fabae, A. solani and A. pisum) did not transmit the

N−Wi PVY isolate. This result confirms previous reports that these aphid species are non to less efficient vectors of PVYN−Wi (Verbeek et al., 2010; Boquel et al., 2011a).

Compared to the other aphids, M. persicae (clone A) showed a significantly higher

PVY transmission efficiency on N. benthamiana (39%). Additionally, the results confirmed that the host plant on which M. persicae is maintained is affecting the

N−Wi transmission efficiency of PVY . The transmission efficiency of PVY was lower when aphids were kept on broad bean plants (50%) compared to Brussels sprouts (81%), with potato and sweet pepper showing intermediate transmission efficiencies (67% and 69%, respectively). This finding is comparable to the findings of Al-Mrabeh (2010) who showed that the transmission efficiency of PVY was low for M. persicae when maintained on potato, as opposed to oilseed rape and tobacco. A possible explanation for these observed differences in PVY transmission, depending on the host plant on which the aphid is reared, could be linked to the constituents of the host plant in

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question. For example, Bosquee et al. (2014) reported that the presence of lectins in plants could interfere with the transmission of PVY and the Cucumber mosaic virus

(CMV). Since broad bean is known to contain lectins (Loewus and Tanner 1982), this could explain the observed 50% transmission efficiency for PVY. Nevertheless, a more detailed study will be needed to confirm this hypothesis for all the plants tested.

So far, the direct molecular quantification of potyviruses has only been conducted in the whole body of single aphids (Singh et al., 1996; Cambra et al., 2004; Olmos et al.,

2005; Moreno et al., 2007) and not in stylets for which reports are limited to detection

(Pelletier et al., 2012; Boquel et al., 2013; Zhang et al., 2013). Hence, this is the first report focusing on virus quantification in the aphid stylet and with attempts to correlate this to the transmission efficiency. Interestingly, a clear correlation was observed between the quantity of PVY in the stylet and the transmission efficiencies of PVY.

While no PVY transcripts were detected in the stylets of the non-vector aphid A. solani, the quantity of PVY in the stylet of M. persicae maintained on broad bean was less than in the stylet of M. persicae on Brussels sprouts. This study confirmed that the quantity of PVY in the stylet is directly correlated with the vector transmission efficiency, and as such could be further evaluated and exploited as a useful index for virus transmission efficiency, which can be used in epidemiological studies later on.

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Chapter 4

The cuticle protein MPCP2 is involved in Potato virus Y transmission in the green peach aphid Myzus persicae

This chapter was published in:

Bahrami Kamangar S, Christiaens O, Taning CN, De Jonghe K, Smagghe G (2019)

The cuticle protein MPCP2 is involved in potato virus Y transmission in the green peach aphid Myzus persicae. Journal of Plant Diseases and Protection, 126: 351-357. https://doi.org/10.1007/s41348-019-00232-w

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Chapter 4

The cuticle protein MPCP2 is involved in Potato virus Y transmission in the green peach aphid Myzus persicae

4.1. Introduction

Potato virus Y (PVY) belongs to the Potyvirus genus and is well known as a destructive and widespread virus that is spread by several aphid vectors in important staple crops such as potato (Wale et al., 2008). Aphids transmit PVY in a non-persistent manner, which implies that acquisition and transmission of virions occur in seconds, when the stylet probes the leaf or just by superficial penetration. The most efficient vector, the green peach aphid Myzus persicae, can transmit PVY with more than 80% efficiency

(Verbeek et al., 2010; Boquel et al., 2011a). Previous studies indicated that Potyvirus particles are retained in the aphid stylet by viral coat proteins (CP) and helper component proteinases (HCPro), both essential molecules in successful Potyvirus transmission (Ammar 1994; Valli et al., 2014). Based on the bridge hypothesis, two domains (KITC and PTK) of the HCPro interact with CP and a proteinaceous receptor

(Pirone and Blanc, 1996) in distal parts of the aphid stylet to retain Potyvirus particles

(Dombrovsky et al., 2007a). A defect in either of the motifs will result in loss of helper function and in non-transmissible viruses (Pirone and Blanc, 1996; Wang et al., 1996;

Martin et al., 1997; Bradley and Ganong, 1955a, b; Harris and Harris, 2001).

However, Harris and Harris (2001) found that transmission of Potyvirus isolates with defects in the HC KITC domain could be rescued in the presence of intact HCs of other related potyviruses.

Identification of receptors of semi- or non-persistent transmissible viruses has been the subject of several studies. Based on in vitro protein–protein interactions, cuticular

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proteins (CuPs) are putative receptors for studied potyviruses with non-persistent transmission (Dombrovsky et al., 2007a). CuP genes from M. persicae, named mpcp2, mpcp5 and mpcp1, interacted with HCPro of Zucchini yellow mosaic virus (ZYMV) and showed 52, 47 and 37% coverage of putative CuP genes, mpcp2 (DQ108935), mpcp5

(DQ108939) and mpcp1 (AF435075), of M. persicae, respectively. However, when these genes and two more CuP genes [including mpcp4 (DQ108938) and mpcp3

(DQ108936)] were expressed in bacteria encoding for CuPs from M. persicae genomes, they did not interact with the HC of ZYMV (Dombrovsky et al., 2007a). These authors suggested that the differences in structure or composition of expressed CuPs in the bacteria and the aphids could be the reason for the lack of interaction of the expressed proteins with HCPro. In a recent study, silencing of the CuP gene mpcp1 in M. persicae by feeding on Arabidopsis producing dsRNA specific for mpcp1 caused a reduction in aphid fecundity by 40 and 47% after 8 and 15 days (Bhatia and

Bhattacharya, 2018). Moreover, ribosomal protein S2 and some other proteins from M. persicae with some similarity to proteins from Buchnera aphidicola (GroEL;

AAR21862), Apis mellifera (hypothetical protein; XP_397538) and Aphis gossypii (CuP;

AAO63549) also interacted with the HC of the potyviruses ZYMV (Dombrovsky et al.,

2007a) and tobacco etch virus (TEV) (Fernandez-Calvino et al., 2010). Receptor localization for the transmission of the NCSP virus, cauliflower mosaic virus (CaMV), has also revealed that the P2 protein (which acts as HCPro in potyviruses) can bind to a proteinaceous compound in the common duct of the distal part of the stylet. This is characterized by a structure named the acrostylet, containing a high concentration of

CuPs (Uzest et al., 2007, 2010). Recently, Webster et al. (2017) revealed that the acrostylet contains a complex structure with available domains of CuPs (mainly from the RR-2 family) interacting with CaMV and another non-persistent transmissible virus

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cucumber mosaic virus (CMV) (Cucumovirus). They identified sequences in the CuPs that could be involved in virus binding. The CuPs belonging to the RR1 or RR2 groups could interact with the HC of Potyvirus or the P2 protein of CaMV (Dombrovsky et al.,

2007a, b; Uzest et al., 2010). Liang and Gao (2017) showed that the expression of certain

CuPs (MPCP1, MPCP2, MPCP4, MPCP5) in M. persicae was increased after feeding on plants which were infected by CMV. Furthermore, they also found that the retention of CMV in M. persicae was reduced when the gene encoding MPCP4 (DQ108938) was knocked down by RNA interference (RNAi) after 48 h of feeding on dsRNA. Further evidence for CuP involvement in virus transmission, via the circulative–propagative transmission pathway, was provided when Liu et al. (2015) discovered that the CPR1

CuP of the plant hopper Laodelphax striatellus interacted with nucleocapsid protein

(pC3) of Rice stripe tenuivirus (RSV). Silencing of this CuP and a yeast-2-hybrid approach showed a decrease in the RSV virion concentration in the haemolymph and salivary glands, as well as a reduction in viral transmission efficiency (Liu et al., 2015).

Cuticle proteins are synthesized in the epidermis or fat body and can also be recycled from old cuticle tissue during ecdysis and metamorphosis stages and reused in the new cuticle (Csikos et al., 1999). These processes are regulated by moulting hormones such as juvenile and ecdysteroid hormones (Riddiford et al., 2003). Cuticle deposition can be controlled by photoperiod shortening and is regulated by the circadian clock in many insects (Ito et al., 2008; Gallot et al., 2010). The largest CuP family in arthropods, the CPR family, is characterized by a chitin-binding domain (CBD) and is divided into

RR-1, RR-2 and RR-3 groups according to three distinct forms of the conserved motif named R and R Consensus. RR-1 is more often associated with the structure of soft or flexible cuticles, while RR-2 is more often found in hard cuticles (Willis et al., 2005). RR genes often cluster in the genome as multi-gene families and are co-regulated

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(Cornman et al., 2008; Togawa et al., 2008). Sequence data from several CuPs of M. persicae and other aphids suggested that they are a typical gene family to aphid species

(Dombrovsky et al., 2003). The role of viral HC and CP proteins in virus transmission has been well studied, yet so far there are no clear in vivo data available on CuPs as the putative receptor molecules in aphids and this not only for PVY, but also for other potyviruses. RNAi as a powerful genetic tool in the study of gene function in different organisms (Gaur et al., 2011), including aphids (Pitino et al., 2011), is increasingly being used.

In this study, we selected five CuPs [MPCP1 (AF435075), MPCP2 (DQ108935),

MPCP3 (DQ108936), MPCP4 (DQ108938) and MPCP5 (DQ108939) that are grouped in RR1 or RR2 CuPr groups] based on their hypothesized involvement in virus transmission in the literature (Dombrovsky et al., 2007a, b; Liang and Gao,

2017). We used an in-house developed feeding system with a liquid artificial diet to deliver gene-specific dsRNA to the aphids and so to silence the gene expression of these CuPs. The post-transcriptional mechanism known as RNA interference (RNAi) is a useful tool to study the functionality of genes of interest. So, with these CuPs exhibiting a reduction in expression, we can investigate their impact on the transmission of the virus in an experimental set-up using potato and tobacco plants,

PVYWilga strain (PVYNWi) and M. persicae aphids. With these results, we hope to shed light on the impact of CuPs in virus transmission, and when successful, it will be the first manuscript, to the best of our knowledge, investigating the function of CuPs in virus transmission under in vivo conditions.

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4.2. Material and Methods

4.2.1. Plants, aphids and virus

M. persicae were taken from a continuous culture at the Laboratory of Agrozoology,

Ghent University. They were reared on Capsicum annum (sweet pepper) at 23 °C,

45% RH and a 16:8 photoperiod. In all of the bioassays, a potato virus Y Wilga

(PVYNWi) isolate (GenBank Accession No. JQ969039), which was collected from a potato farm in Belgium (Bahrami Kamangar et al., 2014), was used. This strain was maintained on 30-60 days old potato plants (Anosta cultivar), grown indoors at room temperature, throughout the study, and the youngest leaflets were used as the virus source in the virus transmission bioassays. The concentration of the virus in these leaves was determined by DAS-ELISA using PVY polyclonal antibody and IgG conjugate according to manufacturer’s instructions (Bioreba, Reinach, Switzerland).

Nicotiana benthamiana seedlings used as indicator plants for aphid transmission tests were grown in a growth chamber (25 °C and 16:8 Light:Dark) and inoculated with the

PVYN-Wi isolate at second leaf stage by M. persicae treated with dsRNA.

4.2.2. Target gene selection and double‑stranded (ds) RNA synthesis

Based on available reports on the interaction of CuP with HC of potyviruses, and the available sequences of CuPs in GenBank, five CuP genes [mpcp1 (AF435075.1), mpcp2 (DQ108935.1), mpcp3 (DQ108936.1), mpcp4 (DQ108938.1) and mpcp5

(DQ108939.1)] were selected as target genes for silencing (Dombrovsky et al., 2007a, b). The correct annotation and phylogenetic relationship of these CuPs and other available CuP sequences in GenBank were determined by a phylogenetic tree using the maximum likelihood method by MEGA7 software (Oxford University, UK).

T7 promoter-tailed primers for dsRNA synthesis (Table 4.1) were designed using

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the E-RNAi Web application (http://www.dkfz.de/signaling/e-rnai3//). Total RNA of M. persicae was extracted from 13 aphids by RNeasy Plant Mini kit (Qiagen, Venlo, the

Netherlands), and cDNA was synthesized with oligo(dt) primer (Invitrogen, Merelbeke,

Belgium) and SuperScript II Reverse Transcriptase (Invitrogen). Target genes were amplified using a mix of cDNA (2 μl), gene-specific T7-tailed primer (Table 4.1) (1 μl),

FastStart Buffer +Mg 10X (5 μl), dNTP mix 10 mM (1 μl), FS Taq DNA Polymerase

5U/μl (0.4 μl) (Invitrogen) and nuclease-free water (37.6 μl). Thermal cycling conditions were as follows: 94 °C for 2 min: 30 s, five cycles (94 °C for 30 s/60 °C for

30 s/72 °C for 30 s), 28 cycles (94 °C for 30 s/65 °C for 30 s/72 °C for 30 s) and 72

°C for 7 min. PCR products were purified by means of the E.Z.N.A. Cycle-Pure Kit

(Omega Bio-Tec, Norcross, GA, USA). The purified PCR amplicons were used to synthesize dsRNA using the MEGA script RNAi Kit (Ambion, Huntingdon, UK), following the manufacturer’s instructions, except for the fact that the final elution of dsRNA from the column was done in nuclease- free water instead of elution solution supplied with the kit.

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Table 4.1. Primers used in this study

Primer name Genbank ID Sequences Application

dsRR1CP1_F DQ108938 AGAGAGAACGGTTACCAGCCGTAa dsRNA synthesis

dsRR1CP1_R AGAACCGCACATAAGACGGAGATa

dsRR1CP2_F DQ108939 AGATCCCAACATACGGCAATTTTa dsRNA synthesis

dsRR1CP2_R AGAACAACCTTGGGTACGACTGGa

dsRR2CP2_F DQ108935 AGAAAATCGTTACACCACCAGCCa dsRNA synthesis

dsRR2CP2_R AGAACCTGGCTTGTTCAACTTCGa

dsRR2CP3_F DQ108936.1 AGAGTCTTACGCACCAAGACCGTa dsRNA synthesis

dsRR2CP3_R AGAAGATTGTTTGGCTGGAGCACa

dsCPUNKN2_F AF435075 AGAAAGCACACAAATACAGCCCCa dsRNA synthesis

dsCPUNKN2_R AGACAGCGTATCCACCTTCCTTCa qRR1CP1_F DQ108938 GAATCCAGAGCCGTCATCTT qPCR qRR1CP1_R CCGTTATCGGTTTGGAAGTT qRR1CP2_F DQ108939 GTCAAGTACTACGCCGACGA qPCR qRR1CP2_R AGGCGATCAATTCCAAAGAC qRR2CP2_F DQ108935 CATCGAAGTTGAACAAGCCA qPCR qRR2CP2_R GGGCACAGCTAGCGTATTCT qRR2CP3_F DQ108936.1 TGCGGCCTATGAACAACCAA qPCR qRR2CP3_R TGATGGAGTGGGAGCTGAAT qCPUNKN2_F AF435075 ACACCGCTGACGACTACAAC qPCR qCPUNKN2_R CTGGTGCAGAGTATGCTGGT q18S_F – ATTCCCAGTAAGCGCGAGTCATCA References Gene in qPCR q18S_R ACTGCGGTCGTTCAATCGGTAGTA References Gene in qPCR MpActF1 – GGTGTCTCACACACAGTGCC References Gene in qPCR MpActR1 CGGCGGTGGTGGTGAAGCTG References Gene in qPCR

dsGFP-F – TACGGCGTGCAGTGCTa dsRNA synthesis

dsGFP-R TGATCGCGCTTCTCGa

aT7 promoter (TAATACGACTCACTATAGGGAGA) tailed primers

4.2.3. Aphid dsRNA feeding bioassay and RNAi

For the feeding RNAi experiments, second-instar M. persicae nymphs were used. For each dsRNA treatment, 60 individual aphids were placed on an artificial diet containing

818 ng/μl of dsRNA and presented through Parafilm in cages as described by Sadeghi et al. (2009). DsRNA targeting green fluorescent protein (GFP) was used as a control treatment. After 48 h, mortality in the treatments was recorded and the alive and active aphids were used for the virus transmission assay. At the same time, treated aphids

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were collected for quantitative real-time PCR (qPCR) analysis, to evaluate the effectivity of the gene silencing.

4.2.4. qPCR expression analysis of CuP genes after dsRNA feeding

Successful gene knockdown after feeding on the CuPs dsRNA was evaluated by qPCR. Total RNA was extracted from four pooled aphids fed on each target gene dsRNA, and dsGFP as control, using the RNeasy Mini kit (Qiagen), and following the manufacturer’s instructions. The RNA extractions were repeated three times (three biological replications), and concentration and purity were assessed using a Nanodrop

ND-1000 system (Thermo Scientific, Zellik, Belgium). The cDNA was synthesized using the Superscript II kit (Invitrogen), following the manufacturer’s instructions. Next, the samples were analysed by qPCR in two technical replications, using the

CFX96TM Real-Time System (Bio-Rad, Nazareth, Belgium). Two reference genes,

18S rRNA (18S) (Bhatia et al., 2012) and actin (Act) (Bass et al., 2013), were utilized in the qPCR experiments to normalize gene expression (Table 4.1). No-template controls (NTC) and no-RT controls (NRT) were also included. Specific primers (Table

4.1) were designed using the Primer3Plus software (http://primer3plus.com/cgi-

GoTaq). First, the specificity of the primer pairs was investigated by melt curve analysis and the efficiency of the PCR reactions was analysed by setting up standard curves for different dilutions of each reaction. Twenty-microlitre reactions were prepared containing 10 μl of GoTaq qPCR Master Mix, 2X (Promega), 2 μl cDNA target, 0.5 μl of each forward and reverse primers and 7 μl of nuclease-free water. The proprietary dye has spectral properties similar to those of SYBR Green I (excitation at 493 nm and emission at 530 nm). Conditions used for amplification were 3 min at 95 °C, followed by 39 cycles of 10 s at 95 °C and 30 s at 56 °C. Data were analysed by the CFX

Manager software (Bio-Rad), after which, an unpaired t test (p = 0.05) was used to

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compare the normalized mRNA levels of each target gene in the respective test samples compared to the control sample, using the SPSS software (IBM, Armonk,

NY, USA).

4.2.5. Virus transmission

For each RNAi treatment and dsGFP control, 12 aphids in three replications (N = 36) were aspirated and starved for 3 h to prompt probing behaviour. Next, the starved aphids were allowed an acquisition access period (AAP) of 5 min on a PVYN-Wi-infected detached potato leaf and were individually transferred to single N. benthamiana seedlings placed in a small insect proof cage. After 24 h, the aphids were killed using pirimicarb (Pirimor, Syngenta). Four weeks later, symptom development in the tobacco plants was assessed and infection rate percentages were evaluated using polyclonal antibody and IgG conjugate (Bioreba, Reinach, Switzerland) by means of

DAS-ELISA and the procedure was done following the manufacturer’s instructions.

The data were subjected to analysis of variance (ANOVA) and Duncan test in a randomized complete block with the SPSS software (IBM, Armonk, NY, USA).

4.3. Results

4.3.1. Successful gene knockdown by dsRNA feeding for two CuPs

Feeding gene-specific dsRNA targeting the mpcp2 and mpcp1 genes to second- instar nymphs for 48 h led to a respective gene-silencing effect of 63 ± 24% and 75 ±

17%, compared to the control (dsGFP) (Fig. 4.1). Feeding dsRNA targeting the other

3 CuPs did not lead to a change in expression for mpcp3, mpcp4 and mpcp5 (Fig. 4.1a– e).

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4.3.2. Knockdown of mpcp2 in M. persicae led to reduced virus transmission

Analysis of the data on transmission of PVY by M. persicae fed on gene-specific dsRNA-containing artificial diet showed a significant difference between dsRNA treatments. The transmission rate of PVY by aphids fed with dsRNA targeting mpcp2

(31 ± 3%) was significantly reduced by 47% with 99.5% certainty (p = 0.005) as compared to the dsGFP- treated aphids (58 ± 5% transmission rate). The transmission rate of aphids fed on dsRNA against mpcp1 was found to be 19% lower (47 ± 6%) than for the dsGFP-fed aphids, but the difference was not so strong with 86% certainty (p =

0.134) (Fig. 4.1f).

Phylogenetic analysis confirmed that all targeted CuPs were correctly annotated and identified. They are separated into two major clades, one containing MPCP5

Fig. 4.1. a–e: CuP target gene knockdown in M. persicae at 48 h post-feeding with dsRNAs targeting CuP genes (mpcp1, mpcp2, mpcp3, mpcp4, mpcp5) and the control (dsGFP-treated group), (mean ± standard error of three biological repeats of four pooled aphids), asterisk (*) indicate a significant difference (p < 0.05). f : Transmission efficiency of PVY by M. persicae treatmented with dsRNAs targeting CuP genes (coefficient variance = 9.67), (mean ± standard error of three biological repeats of 12 N. benthamiana seedlings) (p < 0.05).

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(DQ108939) with an RR1-motif and another containing MPCP1 (AF435075), MPCP2

(DQ108935), MPCP3 (DQ108936) with an RR2-motif and MPCP4 (DQ108938) with an RR1- motif. The phylogenetic analysis also indicated that although both successfully silenced CuPs contain RR2 motifs, they belong to two separated RR2 subclades in the phylogenetic tree, indicating their low sequence similarity (Fig. 2).

Fig. 4.2. Molecular phylogenetic analysis of selected nucleic acid sequences of CuPs of M. persicae [mpcp1 (AF435075.1), mpcp2 (DQ108935.1), mpcp3 (DQ108936.1), mpcp4 (DQ108938.1), mpcp5 (DQ108939.1)] in GenBank by maximum likelihood algorithm; †Interactions with helper component have been reported by Dombrovsky et al. (2007a, b), *CuPs that were silenced in this study, #CuPs that could not be silenced in this study. Evolutionary analyses were conducted in MEGA7 and include a bootstrap analysis (1000 replicates)

4.4. Discussion

In addition to the structural importance, CuPs are also involved in virus transmission, not only in a non- and semi-persistent manner but also in circulative–propagative manner. They have been reported to interact with HC, P2 proteins or directly with CP

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of non/semi-persistent viruses (Dombrovsky et al., 2007a; Liang and Gao, 2017;

Webster et al., 2017; Bhatia and Bhattacharya, 2018) and nucleocapsid protein of circulative–propagative viruses (Liu et al., 2015). Similar to other close relatives in the

Potyvirus genus (ZYMV and TEV), PVY is transmitted in a non-persistent manner and

‘in vitro’ findings indicate that the HCPro bridges the CPs and CuPs to retain viruses in the distal part of stylet of aphids (Dombrovsky et al., 2007a, b; Verbeek et al., 2010;

Boquel et al., 2011). The HCPro of some potyviruses facilitates transmission of other potyviruses, and this is an indication that the function of CuPs as receptors can also be a shared trait in these potyviruses. Moreover, the CuP gene family in aphids share similarities and exhibit some differences (Dombrovsky et al., 2003) that could be the reason for transmission specificity.

In this study, two RR-2 CuPs were successfully silenced in M. persicae. Silencing of the expression of this single CuP, mpcp2 reduced the PVY transmission already strongly (47%) and with a high certainty (99.5%). This CuP, together with other proteins from M. persicae, had already been reported to interact with the HC of the potyviruses ZYMV (Dombrovsky et al., 2007a) and TEV (Fernandez-Calvino et al.,

2010). This study now also demonstrated the involvement of this CuP in PVY transmission in an in vivo assay. RR-2 CuPs in A. pisum are also important for transmission of CaMV and CMV in the acrostylet in the common duct at the distal part of stylets of A. pisum (Uzest et al., 2010; Webster et al., 2017). The common duct, as a rich RR-2 CuP anatomic part (Webster et al., 2017), is also the most efficient part for PVY transmission (Ammar, 1994). The results presented in this study are consistent with the findings of Liang and Gao (2017), who found that the expression of some CuPs, including MPCP2, was increased by feeding on a CMV-infected plant, and they assumed this could suggest involvement of these CuPs in CMV transmission.

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Moreover, they silenced the mpcp4 and showed that this RR-1 CuP is involved in

CMV transmission. This study together with the results of Liang and Gao (2017) shows that at least two CuPs (MPCP4 and MPCP2) are involved in the transmission of non- transmissible viruses, CMV and PVY, respectively, in M. persicae. CMV presents affinity for MPCP4 but PVY is probably transmitted via MPCP2. However, since only

50% of PVY transmission could be explained by MPCP2, we assume that other CuPs might also be involved in this process. We have at least three CuPs in this study that were not silenced and they could also have an effect in virus transmission.

In this study, targeting mpcp1 by RNAi resulted in a higher gene-silencing efficiency

(75%) as compared to mpcp2 (63%). However, looking at the virus transmission data, we see a lower impact of the gene-silencing effect compared to mpcp2. Indeed, with dsmpcp2-fed aphids, the transmission of PVY was reduced by 47% compared to the control (dsGFP-fed aphids). However, with dsMPCP1, the reduction in PVY transmission was smaller with only 19%. The weak(er) involvement of MPCP1 in PVY transmission may just imply that this CuP is not as relevant for the transmission of this virus. Here, we want to remark that it is also possible that this CuP is low in concentration, or nearly not present, in the transmission hot spot of the distal part of the stylet in M. persicae. Another potential explanation could be that MPCP1 has a much longer protein half-life, leading to an ineffective gene silencing at the protein level. Further research should shed light on this. In conclusion, with the successful

RNAi bioassays that we could perform, our data with aphids in vivo on plants confirm earlier data from in vitro studies which indicated that these CuPs are involved in binding and non-persistent transmission of viruses. Therefore, this study showed to the best of our knowledge for the first time, under in vivo conditions the significant involvement of MPCP2 in the transmission of PVY by M. persicae. This is the first report of a

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silenced cuticle protein gene that reduces PVY and Potyvirus transmission in aphids, and we believe that this concept using RNAi can further be exploited to build strategies for managing the transmission of important viruses by vectors such as aphids.

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Chapter 5

General discussion and future perspectives

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Chapter 5

General discussion and future perspectives

5.1. Variability of PVY strains in Belgium

PVY is among the top 5 most economically important plant viruses that can infect a range of host plants, predominantly Solanaceae, worldwide. The NCNP transmission system by aphids and +ssRNA virus genome facilitates a rapid virus transmission, a big genetic diversity and strain variability (Elena and Sanjuan, 2005; Domingo et al,

2008; Tromas and Elena, 2010). The virus, which has its origins in South America, has diversified in Europe (Fuentes et al., 2019) through mutations, genome fragment exchange (recombination events), and cultivar variability, as well as their interaction

(breaking down resistance and adapting to different cultivars) to create different strains.

This diversity in PVY is classified based on the serological properties, biologic response (symptom development) on standard potato cultivars and host range

(biologic division or grouping), in combination with the genomic sequence and recombination pattern (molecular classification). Mutation and recombination produce

PVY variation, and the prevalent variants are determined by fitness ability of the variants. Resistance of Potato cultivars play a major role in the selection, fitness and prevalence of the new PVY variants in any part of the world, as the O and N strains have been replaced by new recombinants (NWi and NTN) in Europe, USA and

Canada due to their potato breeding programs.

Because of PVY-host interaction (potato cultivars, and PVY genetic diversity), some

PVY strains are more common in some parts of the world than in others. Although in recent years, the recombinant NWi and NTN strains are more important in most parts of Europe, USA and Canada, respectively. This was not always the case since they

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gradually replaced the classic O and N strains over the years.

Our findings in this study showed that the average contamination of seed potatoes has slightly increased from 2.4% to 3.6% compared to 2007 (Rolot, 2009). Rolot (2009) showed that the group strain N has been predominant in Belgian potato fields, and we have confirmed this more accurately as the NTN strain (Bahrami Kamangar et al.,

2014). Although NTN was confirmed as the predominant strain in the country, tuber necrosis symptoms were observed in one of the collected samples, which eventually turned out to be negative for PVY infection. The observed necrosis in this PVY- negative sample may have just been caused by adverse weather conditions, since host interaction and environmental conditions such as temperature are also factors that can induce necrosis (Le Romancer and Nedellec 1997). Moreover, other viruses such as tobacco rattle virus (David et al., 2010) and potato mop-top virus (Stevenson et al., 2009), as well as Verticillium wilt and abiotic stress can equally induce different types of tuber necrosis (Stevenson et al., 2009).

Non-recombinant strains (N and O) in Europe (Green at al., 2020) have changed to

‘early’ recombinant strains due to the occurrence of PVY strains co-infecting the same plant, increasing the chances of genomic fragments to be exchanged, and thus recombination events to take place (Chikh ali et al, 2010a; Chikh ali et al, 2010b). Such recombinant events are a continuous process, with primary recombinant strains being capable of further exchanging genomic fragments with non-recombinant strains. This implies that the occurrence of co-infections of two or more PVY strains in host, which occurred in the 2.5% of the tuber samples that were analysed in 2014 in Belgium, can further form secondary recombinants. This phenomenom has been reported in recent studies where new strains have emerged from fragments of primary recombinant strains, and new recombinants have been produced from older recombinants (Kehoe

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and Jones, 2015; Green et al., 2017; Fuentes et al., 2019). These new recombinant strains could have serious implications to plant health and could present new challenges for PVY management.

In addition to the diversity among PVY strains in Belgium, our phylogenetic study based on the sequences of individual isolates also showed differences among the isolates within each strain. This implies that variability within a PVY strain also needs to be taken into account. Three isolates from the Wilga strain (NWi) that were collected in this study clustered in separate (sub-)branches. One was more closely related to the American group, one to the French group, and the third to the German/Hungarian

NWi isolates (Glais et al., 2002, Nie and Singh, 2003b). In addition, the strain O isolates were more similar to European and Japanese isolates, and the retrieved NTN isolates clustered closer to NTN(A) (Chikh Ali et al., 2007b, 2010b), although subdivided into three branches, indicating genetic diversity and point mutations in the isolates. Finally, the N strain isolates were similar to European N isolates. This information is relevant in the context of cultivar selection against PVY strains/isolates in different regions.

Nowadays, globalisation of seed and germplasm exchange is inevitable and while having various benefits, such as providing the best available genetic resources worldwide, it is also likely to increase the genetic diversity of pathogens and the emergence of new strains of viral agents. This can render PVY management more challenging. For example, when looking at the Belgian NTN and NWi isolates, some of them are closely related to isolates from Europe, yet others are closely related to isolates from the United States.

The continuous increase in strain variability could further complicate PVY management and also PVY diagnostics. PVY detection and strain differentiation relies

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on well-designed primers. The profile of designed primers by Chikh Ali (2010a) for multiplex RT-PCR worked well and differentiated PVY strains in this study, yet it is possible that over time, desired fragments would not amplify anymore as shown in our study due to more point mutations in annealing sites. Failing to detect the pathogen following diversification is a constant threat. However, in the last years, the use of high- throughput sequencing (HTS) techniques in plant virus diagnostics has made it easier to reconstitute full PVY genomes in the samples. This has in turn allowed us to better evaluate the current point of care diagnostics by making adjustments or developing new tests when necessary (Massart et al., 2014; Adams et al., 2018; Maree et al.,

2018). According to our research, NTN is the dominant strain in Belgium, whereas

NWi is a new strain. In a three-years sampling (2009-2011), Bosquée et al. (2016) verified our findings about NTN, but they did not find the NWi strain in Belgium. We anticipated PTNRD in our samples because NTN was the dominant strain, but this symptom was extremely rare, and the only sample with PTNRD, was not PVY contaminated. Spunta is the most PVYNTN-infected cultivar in Belgium (21 from 54 sampled farms) (Table 2.2) that did not display PTNRD. Le Romancer and Nedellec

(1997) proved the presence of extreme resistance genes (Ry) in Spunta cultivar that prevent PTNRD. Furthermore, we found that another PVYNTN-infected cultivar (Bintje) did not exhibit PTNRD in our samples, which is consistent with the findings of Le

Romancer and Nedellec (1997), who detected no or a very low rate (5.9 percent) of

PTNRD on this cultivar in farm conditions. In general, PTNRD is an interaction of the environmental conditions and the genetic diversity of potatoes and the virus, and when the cultivars have resistance genes, the probability of PVYNTN incidence is low. In our research, PVYNTN-contaminated samples were cultivars with varying degrees of PVY resistance, and it is logical that there was no PTNRD (Tab 5.1).

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Table 5.1. Resistance of the cultivars that were found infected with PVYNTN and showed no PTNRD (retrived from https://www.europotato.org/)

Cultivar infected with PVYNTN Resistance to potato virus Y Kennebec Medium to very high Cara High to very high Paramount Low Anosta High to very high Eersteling Low to medium Bintje Low - high Diamant Medium to high Spunta medium to very high

5.2. PVY transmission efficiency by different aphid species

Different aphid species are known to transmit potyviruses (including PVY) (Lacomme et al., 2017). However, the virus transmission efficiency is not the same between these species. The list of aphid species found in Belgian potato farms varies year to year

(Vandereycken et al., 2015; Bosquée et al., 2016), including the transmission efficiency of these species. The majority of these aphid species do not colonize potatoes. This diversity may be attributed to the potato farms' surrounding crops as well as the winter weather conditions. PVY transmission efficiency can even be variable between clones of the same species. As noted in this study, PVY is transmitted by the green peach aphid in the range of 0 to 80%, which confirms the variability in PVY transmission within this species. Differences in virus transmission efficiency by aphids can originate from multiple factors, including environmental conditions such as temperature and humidity, feeding behaviour, host plants and virus receptors in aphids (reviewed by Lacomme et al., 2017). Our results showed that M. persicae is the most efficient aphid in PVY transmission compared to other three tested aphids (A. solani, A. fabae and A. pisum), which have been reported as poor or

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non-vectors (DiFonzo et al., 1997; van Hoof, 1980; Basky and Almasi, 2005; Kanavaki et al., 2006; Verbeek et al., 2010; Boquel et al., 2011; Fox et al., 2017). Identification of efficient vectors of destructive potato viruses such as PVY in the potato farms could help to make right decisions in pest management and pesticide application.

Although M. persicae is the most efficient aphid in PVY transmission, the majority of aphid species that transmit PVY in Belgian potato fields are non-colonizers

(Vandereycken et al., 2015; Bosquee et al., 2016).

5.3. The host plant on which an aphid develops affect its PVY transmission efficiency and PVY quantity

Effects of plants as host of virus or aphids is an important part of transmission story.

The PVY load is not the same in different plant species, and each aphid species prefers different host plants. Moreover, the plants species on which the aphids develop before PVY acquisition is another variable that affects virus transmission which has not been studied well. By means of biological methods (transmission to test plant) and molecular quantitative assay (qPCR), we showed that feeding of M. persicae on different plants, before PVY acquisition, affects its PVY transmission efficiency indirectly. Our results showed a direct correlation between transmission efficiency and virus content in the stylet of M. persicae when they were reared on broad bean compare to Brussels sprouts, potato and sweet pepper. Broad bean is a source of plant lectins (Loewus and Tanner, 1982) and lectins were reported to interfere with

CMV transmission (Bosquee et al., 2014; Francis et al., 2020). Lectins are one of the natural substances encoded in all types of life including plants that are used in plant pest and virus vector management (Loris, 2002; reviewed in Hikal et al., 2017). They could prevent the virus transmission by interrupting virus attachment to receptors of

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vectors (Loris, 2002), destabilizing the virus particles (Mitchell et al., 2017) or by affecting the behaviour of virus vectors (Sprawka and Goławska, 2009). The effect of pure extracted lectins on a number of animal and plant viruses has been reported

(Loewus and Tanner, 1982; Loris, 2002; Balzarini et al., 2004; Killiny et al., 2011;

Hivrale and Ingale, 2013; Bosquee et al., 2014; Mitchell et al., 2017; Francis et al.,

2020), but plants as a feeding source of lectin to reduce transmission has been less studied. In the limited studies, turnip plants that were used to rear M. persicae caused a reduction in the transmission of PLRV (a circulative persistent Luteovirus) (Pinheiro et al., 2016). In addition, ZYMV transmission decreased when M. persicae was reared on mustard (Symmes and Perring, 2007).

In the same way, our results indicated that plants such as broad bean could reduce

PVY transmission. Moreover, the amount of PVY in the stylet of M. persicae was less than that of those reared on the other plants. Previous studies have shown that virus detection in the stylet (as an effective organ in PVY transmission), instead of whole body, is more reliable to determine aphid vectors of NCNP viruses (Pelletier et al.,

2012; Boquel et al., 2013; Zhang et al., 2013). The current study is the first to quantify the virus concentration in the aphid stylet and demonstrated the presence of up to

4600 PVY virion copies in the stylet. We also demonstrated a positive correlation between the PVY quantity in the stylet and virus transmission efficiency. In addition, our results also indicated that there was no correlation between the amount of virus in the whole body and the transmission efficiency of PVY by aphids. The reason and mechanism of transmission reduction still needs to be further investigated. However, we found less virus in the stylets of aphids with less transmission efficiency that were fed on the broad bean. Since aphids fed on broad bean before acquiring the virus, it's more likely that plant material (perhaps lectins) interferes with receptors than the virus

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itself. The virus would be unable to bind to the receptors if they are blocked. Plants that reduced virus transmission caused less virus accumulation in the stylet of aphids, and in non-vector aphids, no virus was detected in the individual stylets.

In an Integrated pest management program for plant viruses, the vector populations are monitored by different types of traps (yellow pan trap). However, quantifying or even detecting viruses in the stylet of trapped vectors can improve the accuracy of virus incidence prediction. PVY quantification in dissected stylets of M. persicae may aid in not only predicting the virus vectors but also determining the more efficient vectors. Since viruliferous vectors are more harmful than non-viruliferous vectors, distinguishing them can help to make better decisions about when and how to use control methods such as pesticide application, which is an economic and an environmental issue. Our study proved that broad bean reduced PVY transmission by

M. persicae. This finding can also be considered in virus transmission management in potato farms to minimize PVY transmission. In such a scenario, the broad bean could be planted as a trap crop, barrier crop, or companion plant in or around the potato farms (Stevenson et al., 2009; Lacomme et al., 2017). The use of a trap crop, a barrier crop, and/or a companion plant is an environmentally friendly practice in the management of virus diseases. We propose broad bean as a useful plant that could be use through the above mentioned scenario to manage PVY transmission in potato fields. Potato fields' proximity to crops that are good aphid hosts increases the likelihood of aphids invading the potato field. Furthermore, aphids reared on brussels sprouts transmit PVY more efficiently than aphids reared on broad bean, potato, or pepper, according to our findings. As a result, planting potatoes near brussels sprout farms, which is a suitable host for M. persicae, should be avoided.

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5.4. Cuticle protein receptors as prospective targets to limit PVY transmission by aphids

The interaction between viral proteins and receptors of NCNP transmitted viruses in aphids, affects virus transmission. The viral proteins of PVY and other potyviruses have been well studied and even the binding amino acid motives of these proteins have been well characterized (Granier et al., 1993; Pirone and Blanc, 1996; Kerlan,

2006; Valli et al., 2017). The receptors of NCNP viruses in aphids have long been confirmed to be localized in the stylet (Ammar et al., 1994; Webster et al., 2018), however, very limited information on the biochemical properties of these receptors is available. Most of the data about these receptors have been collected indirectly (in vitro) using protein-protein interaction or EM visualisation but not through bioassays that directly indicate the role/link of cuticle proteins as receptors of NCNP transmitted viruses. Preliminary molecular studies to identify these receptors by Dembrovsky et al. (2007a, b) and Uzest et al. (2007, 2010) showed that the cuticle protein in aphids could be the receptor of NCNP viruses and binds to HC of these viruses. Uzest et al.

(2010) found that the acrostylet in the common duct of the aphid’s stylet is composed of RR1 and RR2 CuPs, and RR2 is the most probable receptor for the NCNP

Caulimovirus (CaMV).

Recently, RNAi has been utilised in receptor detection of NCNP aphid-transmitted plant viruses (Liu et al., 2015; Webster et al., 2017, 2018; Deshoux et al., 2020).

Silencing of predicted CuPs receptors by means of RNAi and cuticle treatment with enzymes were used to determine potential receptors for the CaMV, the NCSP transmissible Caulimovirus in the cuticle stylet, and it indicated that RR1s are more probable than RR2s CuPs (Deshoux et al., 2020). Webster et al. (2017, 2018) showed that the acrostylet contains a complex structure containing different available domains

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of CuPs (mainly from the RR-2 family). Even so, they showed accessibility of different regions but a few amino acids of the RR-2 CuP domains are on the surface of the acrostyle. One of these RR-2 CuP motifs interacted with CMV and hence could be accessible for CMV at the surface of the acrostyle, however, they did not interacted with CaMV unless treated with chitinase (Liu et al., 2015). The chitinase treatment increased the accessibility and interaction of RR2-antibodies, that indicating submerging states of RR2 CuPs in cuticle of acrostyle. In a separate research CMV transmission reduced when a RR1 CuP was silenced using RNAi (Liang and Gao,

2017) that is a contradictory finding compared to the results of Webster et al. (2017,

2018). Moreover, the three RR2 and only one RR1 CuPs interacted with HC of ZYMV showed by Dombrovsky et al. (2007a, b) earlier. We also silenced two cuticle protein genes using RNAi (mpcp2 (DQ108935) and mpcp1 (AF435075.1), one of them the mpcp2 (RR2) significantly reduced the transmission of the PVY virus. This indicates, for the first time that this RR2 CuP is involved in PVY transmission by M. persicae and could be a receptor candidate (Bahrami Kamangar et al., 2019). Our findings are compatible with previous data by Dombrovsky (2007a, b), who demonstrated three

RR2 CuP interacting HC of another Potyvirus ZYMV in vitro. These results indicate that the receptors of CMV and CaMV, ZYMV and PVY (belong to three separate virus family) could be distinct. Moreover the stylets treated with chitinases interacted more strongly with CaMV (Uzest et al., 2010; Webster et al., 2017, 2018; Deshoux et al.,

2020) that there is more than one interacting motif or array in the RR1 and/or that more receptors (RR2) could be present, than RR1. On the other hand, CMV, ZYMV, PVY and, CaMV all non circulative, NCNP or NCSP transmissible viruses, interact with

CuPs by coat protein, P3 protein and helper component that do not share common sequences. Even the two HC of potyviruses (PVY and ZYMV) show variability in the

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interaction domains of CuP, being the KITC motif in PVY and the KLSC motif in ZYMV

(Granier et al., 1993).

Multi-gene families like RR CuPs (Cornman et al., 2008; Togawa et al., 2008) could express CuP copies sharing major similarity and minor differences. There are many homologs for mpcp2 in different species of aphids (Domrovsky et al., 2007a; Uzest et al., 2007; Bahrami Kamangar et al., 2019; Giordano et al., 2020) but they show some differences in amino acid sequencing. The NCNP transmission system is not highly specific due to these major similarities and minor differences (Pirone and Harris,

1977), and an aphid could transmit many, but not all NCNP viruses.

RNAi, as a tool to study biological pathways, successfully revealed relevance between

CuPs (gene of interest) and PVY transmission (related phenotype) in this research.

Two CuP genes were successfully silenced, however, attempts to silence other CuPs failed. There are many possible hypotheses to explain why not all CuPs could be silenced, some of which include: dsRNA instability, low concentration of dsRNAs, and mRNA inaccessibility due to its secondary structure (Shao et al., 2007). Furthermore, the time point of CuP synthesis, as structural proteins, in aphid nymphs is crucial, and silencing can fail if they are already exist in the aphid nymphs. Further studies will be needed to determine the exact cause of failed silencing. Nevertheless, RNAi successfully demonstrated that CuP genes function in PVY transmission through the silencing of 2 out of the 5 CuP genes targeted in this study. RNAi application to reduce or prevent PVY spread by aphids can also be a useful method in PVY management.

In fact, silencing CuPs can also reduce the aphid population as reported by Liang et al. (2017) and Shang et al. (2019).

Pesticides as a source of environmental pollution and health risk have been significantly reduced in IPM programs and are being substituted by safe alternatives.

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Information on PVY biology and epidemiology is knowledge that will help to optimize the use of IPM toolbox options. Our findings revealed some obscurities in diversity and biology of PVY that could be useful in PVY management through potato breeding programs and vector monitoring in farms. Moreover, based on our results, since broad bean decreases PVY transmission, it can be planted as a trap crop or used in companion planting or multiple cropping system with potato. Although significant advances have been made in PVY research through receptor identification and virus quantification with new molecular techniques, there still remain some knowledge gaps in NCNP virus transmission that warrant further investigation.

5.5. Future recommended work

Based on the findings from the research presented in this thesis, it is recommended that future work should concentrate on the following four innovative questions:

 Do inhibitors bind to the receptors or to virus transmission factors like CP or CuP,

or do the aphids just change feeding behavior? The application of EPG in the

treated aphid with proposed inhibitors will provide an answer of this issue.

 How are the virions of NCNP viruses released from aphid receptors during aphid

feeding?

 How many virus copies are released by an aphid during probing, and are there any

differences in quantities of released virus between untreated and treated aphids

with inhibitors?

 In the aphid stylet, how many different types of receptor are present, and which

ones are involved in virus transmission?

 Why weren't three CuP genes silenced in our experiment? More work should be

done to suppress these non-silenced CuP genes using new primers and/or

evaluating different time points for silencing these genes. Furthermore, new

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genome editing tools such as CRISPR/Cas, could be used to knockout these genes.

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Summary

Potato virus Y (PVY), is a plant virus causing important damage in a number of important crops, particularly in potato. Several PVY strains have been differentiated, all of them causing different symptoms and disease levels on numerous commercial potato cultivars. In Belgium, strains belonging to the N group had been reported as the most prevalent, but no detailed information on the relative importance of the PVY strains in

Belgium was published.

PVY is transmitted in a non-persistent manner by many aphid species and the green peach aphid (Myzus persicae) is the most efficient known vector. Aphid species, host plants and environmental conditions are the main factors that influence transmission efficiency. In addition to these main factors, the host plants that are used to rear the aphids prior to the virus acquisition, can affect the PVY transmission efficiency and this aspect has been paid less attention. Furthermore, the quantification of potyviruses in individual vectors has mainly focused on the whole aphid body and not the stylet alone, whereas an even more precise indicator that could correlate with vector efficiency in non-persistently transmitted viruses is the quantity of the virus in the stylet.

Variation in virus transmission by aphid species is another essential question that needs to be better understood. The important role of the distal part of the aphid’s stylet in PVY transmission is known, but the molecular aspects still need to be further elucidated. PVY transmission relays on receptors and aphid-virus protein interaction and understanding the molecular components that are involved, would provide useful opportunities for virus management. PVY receptors have been localized in the tip of the stylet, however, the nature of the receptors is in obscurity. Cuticle proteins (CuP), being the most likely receptors in non/semi-persistent virus transmission, including

PVY transmission, need to be characterized much better. Based on the above-

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mentioned gaps, this project aimed to provide and elucidate the relative importance of the PVY strains in Belgium, the correlation between host plants in aphid rearing before

PVY acquisition, as well as the transmission efficiency, and finally, the relevance of the

PVY concentration in the stylet of aphid vectors related to the transmission efficiency.

We also evaluated the impact of five CuPs on the transmission of PVY by M. persicae using an RNA interference (RNAi) gene-silencing assay.

The PVY survey in Belgian seed potatoes was performed in 2700 individual tubers from

54 seed potato lots originating from 54 production units in 2010. The results revealed a high PVY incidence and substantial strain diversity in some farms. The dominance of the N group in Belgian seed potatoes was confirmed, while the O strain was only found in a few locations. Further characterization using a multiplex PCR identified 75% of the isolates as belonging to the NTN strain group and surprisingly, 7.5% as the mild Wilga strain (N-Wi). The presence of the N-Wi strain was confirmed and further characterized for the first time in Belgian seed potato production.

When evaluating the correlation between the host and PVY quantity, the study revealed that M. persicae, when reared on broad bean and then exposed to acquire

PVY, transmitted PVY less efficiently (50%), compared to when the aphid was reared on Brussels sprout (81%). Additionally, the transmission percentage was directly correlated with the virus concentration in the stylet (r = 0.7), yet not with the concentration in the intact whole aphid body (r = 0.06). Hence, the virus quantity in the stylet could be exploited as a useful index for virus transmission efficiency in epidemiological studies or predictive schemes.

Using an oral RNAi bioassay where the aphids pierce-suck in an artificial diet that was supplemented with gene-specific dsRNA, the expression of two CuPs, mpcp2

(DQ108935) and mpcp1 (AF435075.1), could be decreased by 63% and 75%,

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respectively. Expression of three other CuPs was not affected. Subsequently, we investigated what the effect is of the RNAi-mediated gene silencing on the transmission of PVY from potato to tobacco plants. These results showed with a high certainty of

99.5%, for the first time in vivo, a significant involvement of MPCP2 with a reduction of 47% (compared to the dsGFP-control) in the transmission of PVY. For MPCP1, the effect was smaller with a reduction of 19% and lower certainty of 86%. These findings as useful information and can be used in PVY epidemiology and management.

Keywords:

Solanum tuberosum, potato virus Y, PVYNTN, PVYN-Wi, virus strain, Potyvirus, aphid,

Myzus persicae, non-persistent transmission, virus quantity, transmission efficiency,

RNAi.

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Samenvatting

Aardappelvirus Y (PVY) is een plantenvirus dat belangrijke schade aanricht in een aantal belangrijke gewassen, en specifiek in de aardappel. Verschillende PVY- stammen zijn aanwezig en ze veroorzaken verschillende symptomen en ziekteniveaus op tal van commerciële aardappelrassen. In België werden stammen die tot de N- groep behoren als de meest voorkomende gerapporteerd, maar er werd tot nu toe geen gedetailleerde informatie over het relatieve belang van de PVY-stammen in

België gepubliceerd.

PVY wordt op een niet-persistente manier overgedragen door veel bladluissoorten en de groene perzikbladluis (Myzus persicae) is de meest efficiënte van de gekende vectoren. De soort bladluis, het waardplanttype en omgevingscondities zijn de belangrijkste factoren die de transmissie-efficiëntie beïnvloeden. Naast deze hoofdfactoren kunnen de waardplanten die worden gebruikt om de bladluizen te kweken voorafgaand aan de virusverwerving, de PVY-transmissie-efficiëntie beïnvloeden, en specifiek aan dit aspect werd tot nu toe veel minder aandacht besteed. Bovendien gebeurt de kwantificering van potyvirussen in individuele vectoren voornamelijk op het hele bladluizenlichaam en niet specifiek alleen op het stilet , zijnde de monddelen van de bladluis en de locatie waar de virusoverdracht plaatsvindt. De virusconcentratie in de stilet zou een meer nauwkeurige indicator van de overdrachtsefficiëntie in niet-persistent overgedragen virussen zijn.

Variatie in de efficiëntie van de virusoverdracht door bladluissoorten is een essentiële vraag die beter begrepen zou moeten kunnen worden in functie van de PVY beheersing. De belangrijke rol van het distale deel van de bladluisstilet bij de overdracht van PVY is bekend, maar de specifieke moleculaire aspecten die hier een

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rol bij spelen zijn nog onbekend. De aard van de PVY-overdracht hangt af van receptoren en de interactie tussen bladluis-virus-eiwitten. Het begrijpen van de moleculaire componenten die erbij betrokken zijn, biedt bijkomende kansen voor het virus management. PVY-receptoren zijn gelokaliseerd in de punt van het stilet, maar de aard van de receptoren is onduidelijk. Cuticula-eiwitten (CuP) als de meest waarschijnlijke receptoren bij niet/semi-persistente virusoverdracht, ook bij overdracht van PVY, moeten veel beter worden gekarakteriseerd. Op basis van de bovengenoemde kennisleemtes, had dit doctoraatsproject tot doel de nodige informatie te bekomen over het relatieve belang van de PVY-stammen in België, de correlatie tussen het gebruik van waardplanten in de kweek van bladluizen vóór de blootstelling aan PVY, en de invloed daarvan op de PVY-transmissie-efficiëntie, alsook de relevantie van de PVY-concentratie in de bladluisstilet op de transmissie- efficiëntie. Ook de impact van vijf CuP’s werd onderzocht op de overdracht van PVY door M. persicae met behulp van een RNA-interferentie (RNAi) gen-silencing-assay.

De PVY-survey in Belgisch aardappelpootgoed werd in 2010 uitgevoerd op 2700 individuele knollen die afkomstig waren van partijen uit 54 bedrijven. De resultaten toonden een hoge PVY-incidentie en een aanzienlijke variëteit aan stammen op sommige bedrijven. De dominantie van de N-groep in Belgisch pootgoed werd bevestigd, terwijl de O-stam slechts op enkele locaties werd aangetroffen. Verdere karakterisering met behulp van multiplex PCR bevestigde dat 75% van de isolaten tot de NTN-groep behoorde, en verrassend, 7,5% werd geïdentificeerd als de Wilga-stam

(N-Wi). De aanwezigheid van deze recente N-Wi-stam werd voor het eerst via moleculaire technieken aangetoond en gekarakteriseerd in de Belgische pootaardappelproductie.

De correlatie tussen de gastheer- en de PVY concentratie gaf aan dat M. persicae,

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wanneer deze gekweekt wordt op tuinboon en vervolgens blootgesteld aan PVY acquisitie, PVY minder efficiënt overgedragen werd (50%) in vergelijking met wanneer de bladluis werd gekweekt op spruiten (81%). Bovendien correleerde het overdrachtspercentage direct met de hoeveelheid virus in het stilet (r = 0,7) en totaal niet met de virusconcentratie in het intacte hele bladluizenlichaam (r = 0,06). De virusconcentratie in de stilet zou kunnen een beter bruikbare index zijn om de virus transmissie efficiëntie in te schatten, in ondersteuning van epidemiologische studies of voorspellende schema's.

Met behulp van een orale RNAi-biotoets waarbij de bladluizen zuigen aan een kunstmatig dieet dat werd aangereikt met gen-specifiek dsRNA, kon de expressie van twee CuP’s, mpcp2 (DQ108935) en mpcp1 (AF435075.1), worden verminderd met respectievelijk 63% en 75%. De expressie van de drie andere CuP’s werd niet beïnvloed. Vervolgens werd onderzocht wat het effect is van de RNAi-gemedieerde genuitschakeling op de overdracht van PVY van aardappel- naar tabaksplanten. Deze resultaten toonden voor het eerst in vivo met een hoge zekerheid (99,5%), een significante betrokkenheid van mpcp2 (reductie van 47% vergeleken met de dsGFP- controle) in de PVY overdracht. Voor mpcp1 was het effect kleiner (reductie van 19%) en een lagere zekerheid (86%). Deze bevindingen bieden nuttige informatie in functie van de epidemiologie en het beheer van PVY.

Kernwoorden:

Solanum tuberosum, aardappelvirus Y, PVYNTN, PVYN-Wi, virusstam, Potyvirus, bladluisvector, Myzus persicae, niet-persistente transmissie, virusconcentratie, overdrachtsefficiëntie, RNAi.

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Curriculum vitae

Personal aspects

First name: Saman

Surname: Bahrami Kamangar

Date/Place of birth: September 21, 1961/ Sanandadj, Kurdistan, Iran

Nationality: Iranian

E-mail address: [email protected]

Tel: +98 918 871 5409

Current work address:

Plant Protection Unit, Department of agricultural and horticultural research

Kurdistan Agricultural and Natural Resources Research and Education Center

P.O. Box: 714. postcode:66169-36311

Sanandaj, Kurdistan, Iran.

Current home address:

No. 9, Moalem valey

Kuy farhangian, Shalman Alley

Postal code: 66179-83474

Sanandaj, Kurdistan, Iran.

Education:

M. Sc., Plant pathology - Shiraz University (Iran), (1995-1999)

B. Sc., Plant protection - Urmia University (Iran), (1987-1991)

PhD student in: Department of Plants and Crops, Faculty of Bioscience Engineering,

Ghent University, Ghent, Belgium, and Plant Sciences Unit, Flanders Research

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Institute for Agriculture, Fisheries (ILVO), Merelbeke, Belgium.

Career:

Researcher in plant pathology, Department of Plant Pest and Diseases Research ,

Agricultural and Natural Resources Researches Center of Kurdistan (since 1992).

Department Head of Plant Pest and Diseases Research, Agricultural and Natural

Resources Researches Center of Kurdistan (2002- 2007).

Publications:

Bahrami Kamangar S, Smagghe G, Maes M, De Jonghe K (2014). Potato virus Y

(PVY) strains in Belgian seed potatoes and first molecular detection of the N-WI

strain. J Plant Dis Prot 121:10-19. https://doi.org/10.1007/bf03356485

Bahrami Kamangar S, Mansour Ghazi M, Magowski W, Smagghe G (2016).

Strawberry mite (Phytonemus pallidus fragariae), a new record of tarsonemid

mites (Acari: Tarsonemidae) in Iran. Persian Journal of Acarology 5.4: 351–354.

Bahrami Kamangar S, Van Vaerenbergh J, Kamangar S, Maes M (2017). First report

of angular leaf spot on strawberry caused by Xanthomonas fragariae in

Iran. Plant Dis 101:1031. doi:10.1094/pdis-11-16-1659-pdn

Bahrami Kamangar S, Christiaens O, Taning CN, De Jonghe K, Smagghe G (2019).

The cuticle protein MPCP2 is involved in potato virus Y transmission in the green

peach aphid Myzus persicae. J Plant Dis Prot 126:351-357.

https://doi.org/10.1007/s41348-019-00232-w

Bahrami Kamangar S, Taning CN, De Jonghe K, Smagghe G (2019). Quantity and

transmission efficiency of an isolate of the potato virus Y–wilga (PVYN−Wi) by

aphid species reared on different host plants. J Plant Dis Prot 126: 529–

534. https://doi.org/10.1007/s41348-019-00266-0

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Presentations at conferences:

Afsharifar A, Kamali M, Harighi B, Bahrami Kamangar S, Roumi V, Izadpanah K

(2009) A preliminary Survey of grapevine viruses in Kurdistan province (West of

Iran) (Poster/Extended abstract),16th Meeting of the International Council for the

study of Virus and Virus- like Diseases of the Grapevine (ICVG XVI), p. 114.

Dijon, France, Aug 31-Sept 4, 2009.

https://icvg.org/data/icvg%202009%20part%20I%20%20%20%20pp%201-

131.pdf

Bahrami Kamangar S, Smagghe G, Maes M, De Jonghe K, (2014) Genetic

characterisation of potato virus y isolates from seed potatoes in belgium.

Proceedings of the 19th Triennial Conference of the European Association for

Potato Research (EAPR 2014). 6-11 July 2014 Brussels. (Abstracts book, Poster

session 253). https://edepot.wur.nl/324674

Bahrami-Kamangar S, De Jonghe K, Kamangar S, Maes M, Smagghe G (2010)

Preliminary survey of potato virus Y (PVY) strains in potato samples from

Kurdistan (Iran). Communications in Agricultural and Applied Biological Sciences

75: 783-788.

Bahrami Kamangar S, Christiaens O, De Jonge K, Smagghe G, (2018) Cuticle

proteins are involved in Potato Virus Y transmission in the green peach aphid

Myzus persicae. 70th International Symposium on Crop Protection, Gent

University, Gent, Belgium, 2018/5/22.

Bahrami-Kamangar S, Christiaens O, De Jonghe K, Smagghe G (2018) Potato Virus

Y transmission and quantitative detection in stilet of aphids species. 70th

International Symposium on Crop Protection Gent University, Gent, Belgium.

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