MOLECULAR BIOLOGY AND EPIDEMIOLOGY OF GRAPEVINE LEAFROLL-

ASSOCIATED VIRUSES

By

BHANU PRIYA DONDA

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

DOCTOR OF PHILOSPHY

WASHINGTON STATE UNIVERSITY Department of Plant Pathology

MAY 2016

© Copyright by BHANU PRIYA DONDA, 2016 All Rights Reserved

THANKS

Bioengineering

MAY 2014

© Copyright by BHANU PRIYA DONDA, 2016 All Rights Reserved

To the Faculty of Washington State University:

The members of the Committee appointed to examine the dissertation of BHANU

PRIYA DONDA find it satisfactory and recommend that it be accepted.

Naidu A. Rayapati, Ph.D., Chair

Dennis A. Johnson, Ph.D.

Duroy A. Navarre, Ph.D.

George J. Vandemark, Ph.D.

Siddarame Gowda, Ph.D.

ii ACKNOWLEDGEMENT

I would like to express my respect and deepest gratitude towards my advisor and mentor,

Dr. Naidu Rayapati. I am truly appreciative of the opportunity to pursue my doctoral degree under his guidance at Washington State University (WSU), a challenging and rewarding experience that I will value the rest of my life. I am thankful to my doctoral committee members:

Dr. Dennis Johnson, Dr. George Vandemark, Dr. Roy Navarre and Dr. Siddarame Gowda for helpful advice, encouragement and guidance.

I would like to thank Dr. Sandya R Kesoju (USDA-IAREC, Prosser, WA) and Dr. Neil

Mc Roberts (University of California, Davis) for their statistical expertise, suggestions and collaborative research on the epidemiology of grapevine leafroll disease. To Dr. Gopinath

Kodetham (University of Hyderabad, Hyderabad, India), thank you for believing in me and encouraging me to go the extra mile. I thank Dr. Sridhar Jarugula (Ohio State University

Agricultural Research and Development Center, Wooster, University of Ohio, Ohio, USA), Dr.

Sudarsana Poojari (Agriculture and Agri-Food Canada, Canada) and Dr. Olfemi Alabi (Texas

A&M University, Texas, USA) for help, guidance and motivation during the initial years of my

Ph.D. at WSU. To all members of Dr. Rayapati, past and current, I enjoyed learning alongside you and am grateful for your assistance and support. I sincerely thank the faculty, staff and students in the WSU Department of Plant Pathology for their help and support. It is my pleasure to thank all the members of WSU-Irrigated Agriculture Research and Extension Centre for their help during my PhD.

I wish to acknowledge the following for supporting my graduate research program:

Department of Plant Pathology, WSU Agricultural Research Center, the Wine Advisory

iii Committee, the Washington Wine Commission, Washington State Grape & Wine Research

Program, Northwest Center for Small Fruits Research, and Altria - Chateau Ste. Michelle Wine

Estates. I sincerely acknowledge the following for scholarships and travel awards: Dr. Walter J

Clore scholarship from Washington Wine Industry Foundation, 2015-16 American Society for

Enology and Viticulture scholarship and Student Travel Award to attend the American Society for Enology and Viticulture annual meeting, June 15-18, 2015, Portland, OR, and Raymond G.

Grogan Student Travel Award from the American Phytopathological Society to attend the

American Phytopathological Society annual meeting, August 1-5, 2015, Pasadena, CA.

I am forever grateful to my parents and my brother for their unconditional love and support, which has always been a source of inspiration for me. Lastly, I would like to thank my husband, Kale Harrison, without his encouragement and support this degree would not have been possible. I thank you for being my anchor.

iv MOLECULAR BIOLOGY AND EPIDEMIOLOGY OF GRAPEVINE LEAFROLL-

ASSOCIATED VIRUSES

Abstract

by Bhanu Priya Donda, Ph.D. Washington State University May 2016

Chair: Naidu A. Rayapati

Studies were conducted on molecular biology and epidemiology of grapevine leafroll- associated viruses infecting wine grape (Vitis vinifera) cultivars in Washington State. In the first objective, the complete genome sequence of two isolates of Grapevine leafroll-associated virus

1 (GLRaV-1, genus: Ampelovirus, family Closteroviridae) was determined to be 18,731 and

18,946 nucleotides. The genome of GLRaV-1 isolates contain nine open reading frames with long 5’ and 3’ non-translated regions (NTRs). The sequence differences in the 5’NTR was used to develop a restriction fragment length polymorphism assay for distinguishing GLRaV-1 variants in vineyards. Northern blot hybridization revealed the presence of three of the eight putative 3' co-terminal subgenomic (sg) RNAs at higher levels in virus infected grapevine samples. The 5’ termini of five sgRNAs were mapped and their leader sequences determined.

The results provided a foundation to further elucidate the comparative molecular biology of grapevine-infecting members of the family Closteroviridae.

In the second objective, the spread of grapevine leafroll disease (GLD) was monitored for several seasons in vineyard blocks planted with three red-berried wine grape cultivars.

v Grapevines exhibiting GLD symptoms in these blocks were tested positive for Grapevine leafroll-associated virus 3 (GLRaV-3, genus: Ampelovirus, family Closteroviridae). The temporal spread of GLD indicated higher number of symptomatic vines in each season compared to previous seasons, suggesting increased incidence of the disease during successive seasons.

The spatial distribution of symptomatic vines in all three blocks indicated a disease gradient in which the highest percentage of symptomatic vines were present in rows closest to old vineyard blocks showing GLD symptoms. Spatial autocorrelation (dependence) analysis indicated random distribution of symptomatic vines during initial years of post-planting suggesting primary spread and clustering of symptomatic vines during subsequent years suggesting secondary spread of

GLD. Sequence analysis of a portion of the heat-shock protein 70 homolog gene encoded by

GLRaV-3 revealed predominance of one of the several genetic variants of the virus in the three vineyard blocks. These results provided for the first time science-based knowledge on nature of the spread of GLD in young vineyards to pursue site-specific disease management strategies under conditions prevailing in Washington State.

vi TABLE OF CONTENTS

Page

ACKNOWLEDGEMENT ...... iii

ABSTRACT…...... v

LIST OF TABLES ...... ix

LIST OF FIGURES ...... x

CHAPTER

1. GENERAL INTRODUCTION...... 1

REFERENCES ...... 18

2. SEQUENCE ANALYSIS OF GRAPEVINE LEAFROLL-ASSOCIATED VIRUS 1

ISOLATES FROM WASHINGTON VINEYARDS ...... 34

ABSTRACT ...... 34

INTRODUCTION ...... 35

MATERIALS AND METHODS ...... 38

RESULTS ...... 48

DISCUSSION ...... 70

ACKNOWLEDGEMENTS ...... 77

AFFLIATION OF CO-AUTHORS ...... 77

AUTHORS' CONTRIBUTIONS ...... 77

REFERENCES ...... 78

2. SPATIO-TEMPORAL SPREAD OF GRAPEVINE LEAFROLL-ASSOCIATED

VIRUS 3 IN WASHINGTON VINEYARDS ...... 87

ABSTRACT ...... 87

INTRODUCTION ...... 88

vii MATERIALS AND METHODS ...... 91

RESULTS ...... 97

DISCUSSION ...... 116

ACKNOWLEDGEMENTS ...... 121

AFFLIATIONS OF CO-AUTHORS ...... 121

AUTHORS' CONTRIBUTIONS ...... 122

REFERENCES ...... 123

3. SUMMARY AND CONCLUSIONS ...... 130

viii LIST OF TABLES

Table 1. List of primers used to amplify overlapping genome segments of GLRaV-1 ...... 46

Table 2. List of primers used to determine the 5’- and 3’-terminal sequences of GLRaV-1 genome using 5’RACE kit ...... 46

Table 3. List of primers used to map the 5’ terminus of the sgRNAs of CP, CPd1, CPd2, p21 and p24 from WA-CH and WA-PN isolates ...... 47

Table 4. Comparison of nucleotide and amino acid sequence identity of ORFs between GLRaV-1 isolates...... 54

Table 5. Comparison of the length of 5’ and 3’ NTRs among viruses in family Closteroviridae ...... 58

Table 6. Restriction fragment length polymorphism (RFLP) analysis of the 5’NTR sequence from various GLRaV-1 isolates in Washington...... 66

Table 7. Characteristics of the sub-genomic RNAs specific to the five ORFs encoded by the genome of WA-CH and WA-PN isolates ...... 69

Table 8. Annual incidence of GLD in the three vineyard blocks ...... 105

Table 9. Parameter estimates for disease incidence in the Cabernet Sauvignon block using Gompertz model...... 106

Table 10. Parameter estimates for disease incidence in the Syrah block using Gompertz model ...... 107

Table 11. Parameter estimates for disease incidence in the Petit Syrah block using Gompertz model...... 108

Table 12. Spatial autocorrelation (dependence) analysis using Moran’s I index for GLD incidence in the Cabernet Sauvignon block ...... 111

Table 13. Spatial autocorrelation (dependence) analysis using Moran’s I index for GLD incidence in the Syrah block ...... 111

ix LIST OF FIGURES

Figure 1: Symptoms of GLD in cv. Merlot (red-berried) and cv. Chardonnay (white-berried)...... 5

Figure 2: Genome map of Grapevine leafroll-associated viruses (GLRaVs) in the family Closteroviridae ...... 9

Figure 3: Washington State map displaying current American Viticultural Areas (AVAs) .... 14

Figure 4. Schematic representation of the strategy used for cloning the complete genome of GLRaV-1...... 40

Figure 5. Comparison of L-protease domain among GLRaVs ...... 51

Figure 6. Comparison of AlkB domain among GLRaVs ...... 52

Figure 7. Nucleotide sequence showing the ORF1a-1b overlap and +1 ribosomal frameshift region in GLRaV-1 ...... 53

Figure 8. Multiple sequence alignment of 5’NTR from GLRaV-1 isolates WA-CH, WA-PN and Canada...... 60

Figure 9. MFOLD predicted secondary structures from the 5’NTR sequence of GLRaV-1 isolates WA-CH and WA-PN ...... 61

Figure 10. Multiple sequence alignment of 3’NTR sequence from GLRaV-1 isolates WA-CH, WA-PN, Australia and Canada isolates ...... 62

Figure 11. MFOLD predicted secondary structures from the 3’NTR sequence of GLRaV-1 isolates WA-CH and WA-PN ...... 63

Figure 12. RT-PCR based RFLP method for detecting GLRaV-1 variants in Washington vineyards ...... 65

Figure 13. Northern blot hybridization of GLRaV-1 RNA using riboprobe specific to p24.... 69

Figure 14. Spatial and temporal spread of GLD in a young Cabernet Sauvignon block ...... 101

Figure 15. Spatial and temporal distribution of GLD in a young Syrah block...... 102

Figure 16. Spatial and temporal distribution of GLD in a young Petit Syrah block...... 103

Figure 17. Correlation between GLD symptoms and the presence of GLRaV-3 ...... 104

x Figure 18. Temporal increase in GLD incidence in Cabernet Sauvignon block during 2008 and 2015 seasons ...... 106

Figure 19. Temporal increase in GLD incidence in the Syrah block during 2008 and 2015 seasons ...... 107

Figure 20. Temporal increase in GLD incidence in the Petit Syrah block during 2012 and 2015 ...... 108

Figure 21. Phylogenetic analysis of GLRaV-3 sequences based on partial nucleotide sequences of the HSP70 gene ...... 112

Figure 22. Pie diagram showing the proportion of GLRaV-3 variants analyzed from samples collected from the three vineyard blocks (cvs. Cabernet Sauvignon, Syrah and Petit Syrah) and adjacent old blocks ...... 115

xi CHAPTER ONE

GENERAL INTRODUCTION

The importance of grapes

Grapevine (genus: Vitis and family: Vitaceae) is one of the most valuable horticultural fruit crop grown in many countries worldwide (Troggio et al., 2008). The archeological records suggest the first documentation of grapevine in Eurasia about 65 million years ago (de Saporta,

1879; Bacilieri et al., 2013). Although the wild progenitor of the grapevine (Vitis vinifera subsp. silvestris) was dioecious, the cultivated grapevine (V. vinifera subsp. vinifera) is more diverse, heterozygous and largely hermaphroditic (Zohary, 1996; Aradhya et al., 2003; McGovern, 2004;

Salmaso et al., 2004; This et al., 2006; Troggio et al., 2008; Di Vecchi-Staraz et al., 2009;

Laucou et al., 2011; Bacilieri et al., 2013). Currently, a wide range of cultivars of V. vinifera L. subsp. vinifera are cultivated throughout the world for different purposes, including wine (70%), table grapes (22%) and raisins (8%) and processed products such as juices, jams and jellies

(www.faostat3.fao.org; Troggio et al., 2008). Based on the color variation in the berry skin, cultivars of V. vinifera are broadly classified into two categories, red- and white-berried cultivars

(He et al., 2010). Red-berried cultivars accumulate anthocyanin pigments in the berry skin.

White-berried cultivars lack the anthocyanin pigments in the berry skin. Molecular studies have shown that mutations in the promoter region of the two transcription-factor genes (VvMYBA1 andVvMYBA2) of the MYB-family that regulate transcriptional expression of the UDP- glucose:flavonoid 3-O-glucosyltransferase (UFGT) gene in the anthocyanin pathway was responsible for the loss of color in white-berried cultivars (Kobayashi et al., 2004; Walker et al.,

2007).

1 Grapevine is the most widely planted fruit crop in the world, covering approximately 7.5 million hectares and producing around 69.1 million metric tons of fruit in 2012 (Mullins et al.,

1992; OIV, 2013). After China (~8.0 million) and Italy (~11.6 million), United States is the third largest grape producing country in the world with over 7.7 million metric tons of grapes produced in 2013 (FAOSTAT, 2013). The annual economic impact of grape industry, encompassing wines, grapes, grape products and grape- and wine-associated enterprises, to the

American economy was estimated to be about $162 billion in 2007 (MKF Research, 2007).

Viruses of grapevines

Grapevines are propagated by vegetative cuttings to maintain clonal integrity and trueness-to-type of the progeny vines (This et al., 2006). Currently, about 75 viruses and virus- like agents have been documented in grapevines across the world (Martelli, 2014). As obligate, sub-cellular pathogens, viruses infecting grapevines have been spread around via vegetative cuttings, budwood canes and rootstocks used for grafting (Martelli, 1993, 2000). Apart from other pathogens, viruses have been recognized as one of the major biotic factors limiting the grape production worldwide (Martelli and Boudon-Padieu, 2006; Oliver and Fuchs, 2011).

Among the virus diseases, grapevine fanleaf degeneration/decline, leafroll, rugose wood, fleck and the recently reported red blotch are considered as economically important to grape production in the US (Hewitt, 1954; Bovey and Martelli, 1986; Martelli and Boudon-Padieu,

2006; Calvi, 2011; Krenz et al., 2012; Martelli, 2014; Naidu et al., 2014; Sudarshana et al.,

2015).

Grapevine leafroll disease

2 Historical Perspectives

Historical records available from the mid-19th century indicated the long existence of grapevine leafroll disease (GLD) in Europe and other parts of Mediterranean Basin and Near

East (Gale, 2002). The potential involvement of graft-transmissible agent(s) in the etiology of

GLD was recognized in the early 20th century (Scheu, 1935). Subsequent advances on GLD revealed the presence of long, filamentous viral particles in symptomatic grapevine samples

(Namba et al., 1979). In subsequent years, serological and molecular studies were conducted to demonstrate the presence of several closteroviruses associated with GLD, collectively designated as Grapevine leafroll-associated viruses (GLRaVs; Gugerli et al., 1984; Rosciglione and Gugerli,

1986; Zee et al., 1987; Hu et al., 1990; Zimmermann et al., 1990; Gugerli and Ramel, 1993;

Choueiri et al., 1996; Martelli et al., 2002, 2012; Alkowni et al., 2004; Maliogka et al., 2009;

Ghanem-Sabanadzovic et al., 2010; Maree et al., 2013; Naidu et al., 2014, 2015). Transmission of some of these GLRaVs by several species of mealybugs (Hemiptera: Pseudococcidae) was reported in the 1980’s (Rosciglione and Gugerli, 1989; Tanne et al., 1989; Engelbrecht and

Kasdorf, 1990; Habili et al., 1995). In addition to mealybugs, soft scales (Hemiptera: Coccidae) were recognized as vectors of some GLRaVs in 1994 (Belli et al., 1994). Since then, several species of Pseudococcid mealybugs (reviewed in Daane et al., 2012) and soft scale insects

(Mahfoudhi et al., 2009; Le Maguet, 2012; Kruger and Douglas, 2013) have been reported as vectors of some GLRaVs (Almeida et al., 2013; Naidu et al., 2014). The current status of research on GLRaVs has recently been reviewed (Naidu et al., 2015).

Impact of GLD

3 Several studies have shown that GLD affects vine vigor, fruit ripening, berry sugars, accumulation of anthocyanin pigments in red-berried cultivars, wine quality and overall crop yields (Goheen, 1970; Bovey et al., 1980; Bovey and Martelli, 1992; Golino et al., 2009 a & b;

Lee and Martin, 2009; Lee et al., 2009; Basso et al., 2010; Komar et al., 2010; Alabi et al.,

2012a). An economic study conducted in Finger Lakes region of New York State, indicated an estimated economic loss of $25,000 to $40,000 per hectare in Cabernet franc vineyards depending on the GLD incidence level, yield reduction and impact on fruit quality (Atallah et al.,

2012). A survey of major California grape-growing regions has estimated an economic loss of

$29,902 to $226,405 per hectare in Cabernet Sauvignon vineyards (Ricketts et al., 2015). A recent economic impact study from Washington State estimated that a wine grape grower can lose up to $20,000 per acre over a 20-year period depending on the extent of reduction in fruit yield and berry sugars in a commercial Merlot vineyard (Naidu and Walsh, 2015). These studies showed that GLD can cause substantial impact to vineyard profitability.

Symptoms

One of the unique features of GLD is distinct symptoms produced in red- and white- berried cultivars of V. vinifera (Fig. 1; Rayapati et al., 2008; Naidu et al., 2014). Red-berried cultivars show foliar symptoms consisting of interveinal reddening of leaves with ‘green’ veins.

In contrast, white-berried cultivars, such as Chardonnay, show mild yellowing or chlorosis of leaves (Fig. 1). These symptoms become apparent on mature leaves at bottom portions of canes around véraison (a transitory phase in the berry development representing the onset of berry ripening; Coombe et al., 1992). Symptoms become more apparent in red-berried cultivars than in white-berried cultivars with advancement of the season. Towards the end of the season,

4 symptomatic leaves show downward rolling of leaf margins in both red- and white-berried cultivars. The red and reddish-purple coloration of symptomatic leaves in red-berried cultivars was shown to be due to upregulation of anthocyanin biosynthetic pathway genes and concomitant accumulation of specific classes of anthocyanin pigments (Gutha et al., 2012). In general, cultivars of V. vinifera exhibit symptoms of GLD and those belonging to other species of the genus Vitis may show latent infections (Kovacs et al., 2001).

Figure 1: Symptoms of GLD in cv. Merlot (red-berried) and cv. Chardonnay (white-berried). (A)

Foliar symptoms such as inter-veinal reddening of leaves in cv. Merlot and (C) mild chlorosis of leaves in cv. Chardonnay. (B and D) Downward rolling of leaf margins in both the cultivars.

(Picture source: Naidu et al., 2014).

5 Viruses associated with GLD

Viruses associated with GLD are designated as GLRaVs (family: Closteroviridae). The flexuous filamentous virions of GLRaVs are 1400-2200 nm long and around 12 nm in width and show unique architecture, characteristic of closteroviruses (Namba et al., 1979; Karasev, 2000).

They are numbered serially as GLRaV-1, -2, -3, etc. (Martelli et al., 2002). Among them,

GLRaV-1, -2, -3, -4, and -7 were recognized as distinct species and GLRaV-5, -6, -9, -Pr, -De and -Car were assigned as genetically divergent variants of GLRaV-4 (Martelli et al., 2012).

GLRaV-1, -3, and -4 and its strains belong to the genus Ampelovirus, whereas GLRaV-2 was assigned to the genus Closterovirus and GLRaV-7 to the genus Velarivirus in the family

Closteroviridae (Al Rwahnih et al., 2012; Martelli et. al., 2012; Martelli, 2014; Naidu et al.,

2015). The RNA sequence of GLRaV-8 was shown to be similar to the V.vinifera genome and thus removed from the membership of the genus Ampelovirus (Bertsch et al., 2009; Martelli et al., 2011). The monopartite, positive sense and single-stranded RNA genome of GLRaVs vary in size from 13.6 to 18.6 kilobases (Zhu et al., 1998; Fazeli and Rezaian, 2000; Ling et al., 2004;

Maree et al., 2008; Maliogka et al., 2009; Jarugula et al., 2010b; Jooste et al., 2010; Ghanem-

Sabanadzovic et al., 2010, 2012; Bester et al., 2012, 2014; Jelkmann et al., 2012; Thompson et al., 2012a; Fei et al., 2013; Velasco et al., 2015). Based on their genome size, grapevine- infecting ampeloviruses are broadly divided into two subgroups, GLRaV-1 and GLRaV-3 having large genome size assigned to subgroup I and GLRaV-4 and its strains with smaller genomes, relative to GLRaV-1 and -3, grouped under subgroup II in the genus Ampelovirus (Martelli et al.,

2012).

In addition to differences in genome size, GLRaVs possess variable number of open reading frames (ORFs, Fig.2) that code for proteins involved in a variety of roles in their life

6 cycle. These ORFs are designated from 5’ to 3’ end of the genome according to the convention established earlier for Beet yellows closterovirus (Agranovsky et al., 1994). The unusually large

ORF towards the 5’ end of the genome of GLRaVs is made up of ORF1a and 1b that encode replication-associated proteins (Naidu et al., 2015). Similar to other closteroviruses, ORF1 is expressed from the virion RNA and translation of ORF1a and 1ab polyproteins occurs upon entry into the host cell (Dolja et al., 2006; Gushchin et al., 2013). The ORF1a encodes a large polyprotein containing papain-like protease domain (one or two depending on the virus), a methyltransferase (MET) domain, and a helicase (HEL) domain. L-protease has catalytic Cys and His residues that are characteristic of papain-like proteases found in positive-stranded RNA viruses (Gorbalenya et al., 1991). L-Protease domains in ORF1a are autocatalytically released and are shown to play some important host-specific functional roles (Peng et al., 2001; Liu et al.,

2009). MET and HEL domains contain the DxxR and GKT motifs that are hallmark of positive- strand RNA viruses (Gorbalenya et al., 1990; Rozanov et al., 1992). Viral-encoded helicases are likely involved in unwinding of the double-stranded RNA intermediates and RNA secondary structure (Goregaoker and Culver, 2003). AlkB domain, belonging to the superfamily of 2OG-

Fe(II) oxygenases (Aravind and Koonin, 2001), is present in the genome of some GLRaVs between MET/HEL domains (Naidu et al., 2015). AlkB homologues are present in many RNA viruses (Bratlie and Drablos, 2005), including some viruses infecting woody perennials (Martelli et al., 2007). Similar to the cellular AlkBs, viral AlkBs were proposed to play a role in RNA demethylation and repair of viral RNA (Aas et al., 2003; Dolja et al., 2006; van den Born et al.,

2008). ORF1b encodes the RNA-dependent RNA polymerase (RdRP) domain involved in viral

RNA replication and expressed via +1 ribosomal frameshift mechanism (Dolja et al., 2006). In

7 general, ORF1a and ORF1b forms the replication gene block (RGB) that make up a large portion of the genome of GLRaVs.

The ORFs located downstream of the RGB are variable in number and arrangement in individual GLRaVs (Fig. 2). In the case of GLRaV-1, -2, -3 and -7, the ~6 kDa putative trans- membrane protein, heat shock protein 70 homolog (HSP70h), HSP90h, coat protein (CP) and minor coat protein (CPm) forms a quintuple gene block. In contrast, GLRaV-4 and its strains lack the CPm (Fig. 2; Dolja et al., 2006; Naidu et al., 2015). The order of CP and CPm is reversed in GLRaV-1, -3, and -7 compared to GLRaV-2 (Naidu et al., 2014). GLRaV-1 has an extra copy of the CPm, believed to be originated via gene duplication and divergence during evolution (Boyko et al., 1992; Napuli et al., 2003). Although the role of these proteins in the life cycle of GLRaVs is not yet studied, their functions are likely to be similar to the analogous proteins encoded by other monopartite closteroviruses, such as Beet yellows virus (BYV) and

Citrus tristeza virus (CTV; Agranovsky et al., 1995; Peremyslov et al., 1999, 2004; Tian et al.,

1999; Satyanarayana et al., 2000, 2004; Alzhanova et al., 2001, 2007; Dolja, 2003; Dawson et al., 2013; Dolja and Koonin, 2013). Like in BVY and CTV, proteins encoded by the 3’ proximal

ORFs are likely to be multifunctional playing key roles in cell-to-cell and systemic movement, virion assembly, insect vector transmission, silencing suppression, and other aspects of compatible host-virus interactions (Prokhnevsky et al., 2002; Reed et al., 2003; Lu et al., 2004;

Chiba et al., 2006; Gouveia et al., 2012; Naidu et al., 2015). Analogous to members of the family

Closteroviridae, proteins encoded by the 3’ proximal ORFs, downstream of the RGB, are expressed via 3’-coterminal subgenomic (sg) RNAs (Hilf et al., 1995; He et al., 1997; Fazeli and

Rezaian, 2000; Jarugula et al., 2010b; Dawson et al., 2013; Dolja and Koonin, 2013).

8

Figure 2: Genome map of Grapevine leafroll-associated viruses (GLRaVs) in the family

Closteroviridae. Schematic representation of the genome map of GLRaV-1

(NC_016509), GLRaV-2 (NC_007448), GLRaV-3 (EU259806), GLRaV-4 (NC_016416), and GLRaV-7 (JN383343). Replication gene block (RGB) and Quintuple gene block (QGB) are shown in boxes with dotted lines and the open reading frames are shown as closed boxes in the genome. Abbreviations: L-Pro, Leader protease; AlkB, AlkB conserved domain; MET, methyl transferase; HEL, RNA helicase; RdRP, RNA-dependent RNA polymerase; HSP70h, heat shock protein-70 homologue; CP, coat protein; CPm, minor coat protein. (Picture source: Naidu et al.,

2015).

9 Diagnosis of GLRaVs

GLRaVs can be detected through biological, serological and molecular assays. Biological or field indexing involves grafting a bud wood from a candidate vine on to a susceptible indicator host and observing for symptom development 2 to 3 years after grafting (Rowhani et al., 2005; Constable et al., 2013). However, the limitation of field indexing is that it is time- consuming, labor-intensive and does not identify the virus species present in the test material. In addition, few viruses such as GLRaV-2 and GLRaV-7 were shown to cause asymptomatic symptoms in grapevine (Pourrahim et al., 2007; Bertazzon et al., 2010; Alkowni et al., 2011; Al

Rwahnih et al., 2012; Jelkmann et al., 2012; Poojari et al., 2013a) and thus are not detectable in biological assays. Therefore, sensitive and robust detection of GLRaVs is critical for studying their epidemiology and improving the sanitary status of grapevine propagation material in grapevine certification programs. Serological assays such as enzyme-linked immunosorbent assay (ELISA) is widely used for routine diagnosis of GLRaVs (Rowhani, 1992; Forsline et al.,

1996; Besse et al., 2009). However, the limitations of ELISA include less sensitivity (inability to detect a virus at low concentrations) and specificity (inability to identify new variants of the virus) of detection of GLRaVs in grapevines (Naidu et al., 2014). Molecular techniques, such as reverse transcription (RT)-PCR (Rowhani et al., 2000; Ling et al., 2001; Constable et al., 2012), quantitative real-time PCR (Osman et al., 2007, 2012; Bertolini et al., 2010; López-Fabuela et al., 2013), RT-loop mediated isothermal amplification (Walsh and Pietersen 2013), microarray

(Engel et al., 2010), macroarray (Thompson et al., 2012b) and recently, next generation sequencing (Al Rwahnih et al., 2009, 2013; Coetzee et al., 2010; Zhang et al., 2011;

Giampetruzzi et al., 2012; Poojari et al., 2013b), were used for the detection of viruses and their

10 strains in grapevines. Although these assays are highly sensitive and robust, molecular techniques require expertise and costly equipment relative to biological and serological methods.

Spread of GLRaVs

GLRaVs are phloem-limited and difficult to transmit from grapevine to grapevine by manual inoculations. The spread of GLRaVs occurs mainly via vegetative propagation, a viticultural practice performed by growers to preserve clonal integrity of agronomically valuable traits in grapevines (Schon et al., 2009). GLRaVs can also be transmitted via the scion and/or rootstock used for grafting. Different species of mealybugs (Hemiptera: Pseudococcidae) and soft scale insects (Hemiptera: Coccidae) are known to spread GLRaV-1, -3, and -4 and its strains in a semi-persistent manner (Golino et al., 2002; Sforza et al., 2003; Hommay et al., 2008; Tsai et al., 2010). In Washington State, grape mealybug (Pseudococcus maritimus Ehrhorn) and

European fruit lecanium scale (Parthenolecanium corni Bouché) were reported as vectors of

GLRaV-3 (Bahder et al., 2013). Among these, Ps. maritimus appears to be a more competent vector of GLRaV-3 in California with 40 - 41% rate of transmission achieved (Golino et al.,

2002). No vector has been reported so far for GLRaV-2, although its transmission by manual inoculations to herbaceous hosts has been demonstrated (Boscia et al., 1995; Goszczynski et al.,

1996). GLRaV-7 has no known insect vector either. However, experimental transmission by parasitic dodder (Cuscuta reflexa and C. europea) to herbaceous hosts has been demonstrated

(Mikona and Jelkmann, 2010). The role of insect vectors in the spread of GLRaV-1 and GLRaV-

4 and its strains in Washington State has not yet been determined. However, reports from other parts of the world suggests that GLRaV-4 and its strains can be transmitted by different species of mealybugs, such as vine mealybug (Planococcus ficus), citrus mealybug (Planococcus citri),

11 longtailed mealybug (Pseudococcus longispinus), grape mealybug (Pseudococcus maritimus), obscure mealybug (Pseudococcus viburni), apple mealybug (Phenacoccus aceris) and soft scale insect, Ceroplastes rusci (Gugerli, 2003; Habili et al., 2003; Sim et al., 2003; Mahfoudhi et al.,

2009; Tsai et al., 2010; Le Maguet et al., 2012; Martelli, 2012).

Epidemiology of leafroll disease

Natural spread of GLD was first reported in vineyards of South Africa (Engelbrecht and

Kasdorf, 1985). The association of mealybugs with the disease spread was shown in greenhouse experiments using Pl.ficus (Engelbrecht and Kasdorf, 1990). Following this study, many reports have indicated the spread of GLD in newly planted vineyards in Australia (Habili et al., 1995;

Habili and Nutter, 1997), South Africa (Pietersen, 2006; Sokolsky et al., 2013), USA (Golino et al., 2008), New Zealand (Charles et al., 2009), Italy (Gribaudo et al., 2009) and France (Le

Maguet et al., 2013). In all these studies, two common patterns of spread was reported: primary spread via planting infected plant material or initiated by alighting first instar viruliferous mealybug vectors and secondary spread between adjacent vines within and across rows via movement of viruliferous mealybugs. Several studies have reported the secondary spread of

GLRaV-3 within a vineyard by different species of mealybugs (Habili et al., 1995; Habili and

Nutter, 1997; Cabaleiro and Segura, 2003, 2006; Charles et al., 2006; Cabaleiro et al., 2008;

Golino et al., 2008). The field spread of vector-transmissible GLRaVs can be influenced by a variety of factors, including the vector feeding behavior, grapevine cultivar, age of grapevine at which virus acquisition or inoculation happens, and presence of GLRaVs in single or mixed infection in a grapevine (Naidu et al., 2014). Vineyard spread can also be influenced by other

12 factors such as regional environment, landscape features and viticultural practices that affect the survival and dispersal of mealybugs (Daane et al., 2012).

Wine grapes in Washington State

Within the United States, Washington State ranks second, after California, in wine grape acreage and production. In 2014, 227,000 tons of wine grapes were produced from the state vineyards (www.washingtonwine.org; NASS, 2015). Of the total wine grapes produced from approximately forty different V. vinifera cultivars, nearly 53% accounts from white-berried and

47% from red-berried cultivars (www.washingtonwine.org). Cabernet Sauvignon and Merlot, among the red-berried cultivars, and Riesling and Chardonnay, among the white-berried cultivars, occupy the large portion of vineyard acreage in the state. The economic impact of

Washington State’s grape and wine industry was estimated to be $4.8 billion in 2013 compared to $3.5 billion in 2009, representing a compound annual growth rate of 8.5% (Washington State office of financial management, 2014; Community attributes Inc., 2015; www.washingtonwine.org). Wine grape cultivars are currently planted in about 50,000 acres in

13 American Viticultural Areas (AVA’s) across the state (Fig. 3). They are listed, in the order of their official recognition (year in parenthesis) by Alcohol and Tobacco Tax and Trade Bureau

(TTB) of the US Treasury department as follows: Yakima Valley (1983), Walla Walla Valley

(1984), Columbia Valley (1984), Puget Sound (1995), Red Mountain (2001), Columbia Gorge

(2004), Horse Heaven Hills (2005), Wahluke Slope (2006), Rattlesnake Hills (2006), Snipes

Mountain (2009), Lake Chelan (2009), Naches Heights (2011) and Ancient Lakes (2012) (Fig. 3, www.washingtonwine.org).

13

Figure 3: Washington State map displaying the current American Viticultural Areas (AVAs).

The map shows the 13 different AVAs; namely, including Yakima Valley, Walla Walla Valley,

Columbia Valley, Puget Sound, Red Mountain, Columbia Gorge, Horse Heaven Hills, Wahluke

Slope, Rattlesnake Hills, Snipes Mountain, Lake Chelan, Naches Heights and Ancient Lakes.

(Source: www.washingtonwine.org. Accessed on September 1, 2015).

14 Current Status of GLRaVs

Since 2005, several viruses and viroids have been documented in Washington vineyards.

They include grapevine leafroll-associated viruses (GLRaV-1, -2, -3, and -4 and its strains

GLRaV-5 and -9), Grapevine red blotch-associated virus (GRBaV), Grapevine virus A (GVA),

Grapevine virus B, Grapevine Virus E, Grapevine rupestris stem pitting-associated virus

(GRSPaV), Grapevine fanleaf virus (GFLV), Tobacco ring spot virus, Grapevine fleck virus and

Grapevine Syrah virus-1 and three viroids (Hop stunt viroid, Grapevine yellow speckle viroid 1 and 2) (Martin et al., 2005; Soule et al., 2006; Naidu, 2011; Alabi et al., 2012b; Bahder et al.,

2013a; Poojari et al., 2013a). Molecular studies have shown the presence of genetically and biologically distinct variants of GLRaV-1 and -2, GRSPaV, GFLV, and GVA in Washington vineyards (Mekuria et al., 2009; Alabi et al., 2010; Jarugula et al., 2010a; Alabi et al., 2011;

Poojari et al., 2013a; Alabi et al., 2014). These viruses can be found as single and/or mixed infections in individual grapevines (Naidu, 2011). GLRaV-3 was found to be the most common and widespread among all the GLRaVs documented in Washington vineyards (Martin et al.,

2005; Naidu, 2011). Grape mealybug (Pseudococcus maritimus Ehrhorn) and European fruit lecanium scale (Parthenolecanium corni Bouché) were reported as vectors of GLRaV-3 (Bahder et al., 2013).

Objectives and justification of the study

Owing to the deleterious effects of viruses to wine grape production, the Washington

State Grape Industry has identified “management of viruses that impact fruit quality and vine health” as one of the highest priorities for “achieving the central goal of tripling the economic value of the wine and juice industry by 2020” in its Research Task Force Report “Building the

15 Future of Washington State Grape and Wine Industry through Research” published in 2008.

Similarly, the Wine Advisory Committee of the Washington State Wine Commission

(www.washingtonwine.org) and the Washington Association of Wine Grape Growers

(WAWGG, www.wawgg.org) have recognized virus diseases as a significant impediment to sustainable growth of the wine grape industry in the state. Among the several virus diseases documented in Washington vineyards, GLD continues to be a significant threat to the productivity of vineyards (Rayapati et al., 2008; Alabi et al., 2012a). Thus, management of GLD is one of the high priorities for Washington’s grape and wine industry. Therefore, science-based knowledge on different aspects of GLD will help in advancing disease management strategies for sustainable growth of this rapidly expanding industry in Washington State.

Previous research related to GLD in Washington vineyards was largely focused on molecular biology and impacts of GLRaV-3 (Jarugula et al., 2010a and b; Alabi et al., 2012a) and genetic diversity of GLRaV-1 and -2 (Jarugula et al., 2010a; Alabi et al., 2011; Poojari et al.,

2013a). To date, complete genome sequence of GLRaV-2, -3, -7, -4, and its strains -6, -9, -Pr and

-Car are available (Zhu et al., 1998; Ling et al., 2004; Maliogka et al., 2009; Ghanem-

Sabanadzovic et al., 2010, 2012; Jelkmann et al., 2012; Velasco et al., 2015). In contrast, only partial genome sequence of GLRaV-1, the second most widely distributed virus across many grapevine-growing regions, is available in the literature (Fazeli and Rezaian, 2000; Habili et al.,

2007). Thus, determining the complete genome sequence of GLRaV-1 would help in examining the comparative molecular biology of GLRaVs. Although studies on the epidemiology of GLD has been conducted in different grapevine-growing regions in the US, South Africa, Australia,

New Zealand and many European countries (Engelbrecht and Kasdorf, 1985; Habili et al., 1995;

Habili and Nutter, 1997; Cabaleiro and Segura, 2003, 2006; Charles et al., 2006, 2009; Pietersen,

16 2006; Cabaleiro et al., 2008; Golino et al., 2008; Cabaleiro, 2009; Gribaudo et al., 2009; Almeida et al., 2013; Le Maguet et al., 2013; Sokolsky et al., 2013), very little research has been conducted on field spread of GLD in Washington vineyards. Although previous studies have indicated that GLRaV-3 is more widespread across Washington vineyards (Martin et al., 2005;

Naidu, 2011), many outstanding questions regarding field spread of GLRaV-3 remains to be answered. Growers have been encouraged to use virus-tested planting stock for planting new vineyard blocks as a first line of defense for minimizing the spread of GLD. However, the risk of

GLD spread into new plantings in time and space needs to be addressed, if appropriate control strategies are to be implemented to mitigate negative impacts of the disease.

This study was undertaken on two specific aspects of GLD as listed below:

(i) Conduct studies on molecular aspects of Grapevine leafroll-associated virus 1 to

determine the complete genome sequence of the virus and examine 3’-coterminal

subgenomic RNAs, and develop a molecular method to distinguish virus variants.

(ii) Conduct studies on the epidemiology of grapevine leafroll disease with emphasis on

the spatial and temporal spread of the disease in young vineyards planted with virus-

tested planting stock.

17 REFERENCES

Aas, P. A., Otterlei, M., Falnes, P. O., Vågbø, C. B., Skorpen, F., Akbari, M., Sundheim, O., Bjørås, M., and Slupphaug, G. 2003. Human and bacterial oxidative demethylases repair alkylation damage in both RNA and DNA. Nature 421:859–863.

Agranovsky, A. A., Koonin, E. V., Boyko, V. P., Maiss, E., Frotschl, R., Lunina, N. A., and Atabekov, J. G. 1994. Beet yellows closterovirus: complete genome struture and identification of a leader papain-like thiol protease. Virology 198:31-324. doi:10.1006/viro.1994.1034.

Agranovsky, A. A., Lesemann, D. E., Maiss, E., Hull, R., and Atabekov, J. G. 1995. “Rattlesnake” structure of a filamentous plant RNA virus built of two capsid proteins. Proc. Natl. Acad. Sci. U.S.A. 92:2470–2473. doi:10.1073/pnas.92.7.2470.

Alabi OJ, Martin RR, Naidu RA. 2010. Sequence diversity, population genetics and potential recombination events in Grapevine rupestris stem pitting-associated virus in Pacific North-West vineyards. J. Gen. Virol. 91:265-276.

Alabi, O. J., Al Rwahnih, M., Karthikeyan, G., Poojari, S., Fuchs, M., Rowhani, A., and Naidu, R. A. 2011. Grapevine leafroll-associated virus 1 occurs as genetically diverse populations. Phytopathol. 101:1446–56. doi:10.1094/PHYTO-04-11-0114.

Alabi, O. J., Gutha, L. R., Casassa, L. F., Harbertson, J., Mirales, M., Beaver, C. W., Davenport, J., and Naidu, R. A. 2012a. Impacts of grapevine leafroll disease on own- rooted wine grape cultivar in cool climate conditions. 63rd American Society for Enology and Viticulture National Conference. Portland, OR, U.S.A, pp. 69-70.

Alabi, O. J., Zheng, Y., Jagadeeswaran, G., Sunkar, R., and Naidu, R. A. 2012b. High- throughput sequence analysis of small RNAs in grapevine (Vitis vinifera L.) affected by grapevine leafroll disease. Mol. Plant Pathol. 13:1060–1076. doi:10.1111/j.1364- 3703.2012.00815.x.

Alkowni, R., Rowhani, A., Daubert, S., and Golino, D. 2004. Partial characterization of a new ampelovirus associated with grapevine leafroll disease. J. Plant Pathol. 86:123–133.

Alkowni, R., Zhang, Y. P., Rowhani, A., Uyemoto, J. K., and Minafra, A. 2011. Biological, molecular, and serological studies of a novel strain of grapevine leafroll-associated virus 2. Virus Genes. 43:102-110. doi:10.1007/s11262-011-0607-7.

Almeida, R. P. P., Daane, K. M., Bell, V. A., Blaisdell, G. K., Cooper, M. L., Herrbach, E., and Pieterson, G. 2013. Ecology and management of grapevine leafroll disease. Front. Microbiol. 4:94. doi:10.3389/fmicb.2013.00094.

Al Rwahnih, M., Daubert, S., Golino, D., and Rowhani, A. 2009. Deep sequencing analysis of RNAs from grapevine showing Syrah decline symptoms reveals a multiple virus infection that includes a novel virus. Virology 387:395-401.

18

Al Rwahnih, M., Dolja, V. V., Daubert, S., Koonin, E. V., and Rowhani, A. 2012. Genomic and biological analysis of grapevine leafroll-associated virus 7 reveals a possible new genus within the family Closteroviridae. Virus Res. 163:302–309. doi:10.1016/j.virusres.2011.10.018.

Al Rwahnih, M., Dave, A., Anderson, M., Rowhani, A., Uyemoto, J. K., and Sudarshana, M. R. 2013. Association of a DNA virus with grapevines affected by red blotch disease in California. Phytopathology 103:1069-1076.

Alzhanova, D. V., Napuli, A. J., Creamer, R., and Dolja, V. V. 2001. Cell-to-cell movement and assembly of a plant closterovirus: roles for the capsid proteins and HSP70 homolog. EMBO J. 20:6997–7007. doi:10.1093/emboj/20.24.6997.

Alzhanova, D. V., Prokhnevsky, A. I., Peremyslov, V. V., and Dolja, V. V. 2007. Virion tails of Beet yellows virus: Coordinated assembly by three structural proteins. Virology 359:220–226.

Aradhya, M. K., Dangl, G. S., Prins, B. H., Boursiquot, J.-M., Walker, M. A., Meredith, C. P., and Simon, C. J. 2003. Genetic structure and differentiation in cultivated grape, Vitis vinifera L. Genet. Res. 81:179–192.

Aravind, L., and Koonin, E. V. 2001. The DNA-repair protein AlkB, EGL-9, and leprecan define new families of 2-oxoglutarate- and iron-dependent dioxygenases. Genome Biol. 2:7.1-7.8.

Atallah, S. S., Gómez, M. I., Fuchs, M. F., and Martinson, T. E. 2012. Economic impact of grapevine leafroll disease on Vitis vinifera cv. Cabernet franc in Finger Lakes vineyards of New York. Am. J. Enol. Vitic. 63:73–79. doi:10.5344/ajev.2011.11055.

Bacilieri, R., Lacombe, T., Le Cunff, L., Di Vecchi-Staraz, M., Laucou, V., Genna, B., Peros, J- P., This, P., and Boursiquot, J-M. 2013. Genetic structure in cultivated grapevines is linked to geography and human selection. BMC Plant Biol. 13:25. doi:10.1186/1471-2229-13-25.

Bahder, B. W., Poojari, S., Alabi, O. J., Naidu, R. A., and Walsh, D. B. 2013. Pseudococcus maritimus (Hemiptera: Pseudococcidae) and Parthenolecanium corni (Hemiptera: Coccidae) are capable of transmitting grapevine leafroll-associated virus 3 between Vitis x labruscana and Vitis vinifera. Environ. Entomol. 42:1292–1298. doi:10.1603/EN13060.

Bertsch, C., Beuve, M., Dolja, V. V., Wirth, M., Pelsy, F., Herrbach, E., and Lemaire, O., 2009. Retention of the virus-derived sequences in the nuclear genome of grapevine as a potential pathway to virus resistance. Biol. Direct. 4:1–11.

Besse, S., Bitterlin, W., and Gugerli, P. 2009. Development of an ELISA for the simultaneous detection of Grapevine leafroll associated virus 4, 5, 6, 7, and 9. Pages 296-298 in: Proc. XVI Int. Counc. Study Viruses VirusLike Dis. Grapevine.

Basso, M. F., Fajardo, T. V. M., Santos, H. P., Guerra, C. C., Ayub, R. A., and Nickel, O. 2010. Leaf physiology and enologic grape quality of virus-infected plants. Trop. Plant Pathol. 35:351–

19 59. doi:10.1590/S1982-56762010000600003.

Belli, G., Fortusini, A., Casati, P., Belli, L., Bianco, P. A., and Prati, S. 1994. Transmission of a grapevine leafroll associated closterovirus by the scale insect Pulvinaria vitis L. Riv. Patol. Veg. 4:105–108.

Bertazzon, N., Borgo, M., Vanin, S., and Angelini, E. 2010. Genetic variability and pathological properties of grapevine leafroll-associated virus 2 isolates. Eur. J. Plant Pathol. 127:185-197. doi:10.1007/s10658-010-9583-3.

Bertolini, E., García, J., Yuste, A., and Olmos, A. 2010. High prevalence of virus in table grape from Spain detected by real-time RT-PCR. Eur. J. Plant Pathol. 128: 283-287.

Bester, R., Maree, H. J., and Burger, J. T. 2012. Complete nucleotide sequence of a new strain of grapevine leafroll-associated virus 3 in South Africa. Arch. Virol.157:1815– 19. doi:10.1007/s00705-012-1333-8.

Bester, R., Pepler, P. T., Burger, J. T., and Maree, H. J. 2014. Relative quantitation goes viral: An RT-qPCR assay for a grapevine virus. J. Virol. Methods 210:67-75. doi:10.1016/j.jviromet.2014.09.022.

Boscia, D., Greif, C., Gugerli, P., Martelli, G.P., Walter, B., and Gonsalves, D. 1995. Nomenclature of grapevine leafroll-associated putative closterovirus. Vitis 34:171-175.

Bovey, R., Gartel, W., Martelli, G.P., and Vuitenez, A. 1980. Virus and Virus-like Diseases of Grapevines. Editions Payot, Lausanne.

Bovey, R., and Martelli, G. 1986. The viroses and virus-like diseases of the grapevine. A bibliographic report, 1979-1984. Vitis 25:227-275.

Bovey, R., and Martelli, G.P., 1992. Directory of major virus and virus-like diseases of grapevines. Mediterranean Fruit Crop Improvement Council (MFCIC), Rome, pp. 41–51.

Boyko, V. P., Karasev, A. V, Agranovsky, A. A., Koonin, E. V., and Dolja, V. V. 1992. Coat protein gene duplication in a filamentous RNA virus of plants. Proc. Natl. Acad. Sci. U.S.A. 89:9156–9160.

Bratlie, M. S., and Drablos, F. 2005. Bioinformatic mapping of AlkB homology domains in viruses. BMC Genomics 6:1. doi:10.1186/1471-2164-6-1.

Cabaleiro, C., and Segura, A. 2003. Monitoring the field spread of Grapevine leafroll associated virus 3 for 12 years. Pages 216-217 in: Proc. XIV Int. Counc. Study Viruses Virus-Like Dis. Grapevine.

Cabaleiro, C., and Segura, A. 2006. Temporal analysis of Grapevine leafroll associated virus 3 epidemics. Eur. J. Plant Pathol. 114:441-446.

20

Cabaleiro, C., Couceiro, C., Pereira, S., Cid, M., Barrasa, M., and Segura, A. 2008. Spatial analysis of Grapevine leafroll associated virus 3 epidemics. Eur. J. Plant Pathol. 121:121-130.

Cabaleiro, C. 2009. Current advances in the epidemiology of grapevine leafroll disease. Pages 264-268 in: Extended Abstr. 16th Meet. ICVG. Dijon, France.

Calvi, B. L. 2011. Effects of red-leaf disease on Cabernet Sauvignon at the Oakville experimental vineyard and mitigation by harvest delay and crop adjustment. MS thesis, University of California, Davis.

Charles, J. G., Cohen, D., Walker, J. T. S., Forgie, S. A., Bell, V. A., and Breen, K. C. 2006. A review of the ecology of grapevine leafroll associated virus type 3 (GLRaV-3). N.Z. Plant Prot. 59:330-337.

Charles, J. G., Froud, K. J., Brink, R. V. D., and Allan, D. J. 2009. Mealybugs and the spread of grapevine leafroll-associated virus 3 (GLRaV-3) in a New Zealand vineyard. Australasian Plant Pathol. 38:576–583.

Chiba, M., Reed, J. C., Prokhnevsky, A. I., Chapman, E. J., Mawassi, M., Koonin, E. V., Carrington, J. C., and Dolja, V. V. 2006. Diverse suppressors of RNA silencing enhance agroinfection by a viral replicon. Virology 346:7–14. doi:10.1016/j.virol.2005.09.068.

Choueiri, E., Boscia, D., Digiaro, M., Castellano, M. A., and Martelli, G.P. 1996. Some properties of a hitherto undescribed filamentous virus of the grapevine. Vitis 35:91–93.

Coetzee, B., Freeborough, M., Maree, H. J., Celton, J., Rees, D. J., and Burger, J. T. 2010. Deep sequencing analysis of viruses infecting grapevines: Virome of a vineyard. Virology 400:157- 163.

Constable, F. E., Connellan, J., Nicholas, P., and Rodon, B. C. 2012. Comparison of enzyme- linked immunosorbent assays and reverse transcription polymerase chain reaction for the reliable detection of Australian grapevine viruses in two climates during three growing seasons. Aust. J. Grape Wine Res. 18:239-244. doi:10.1111/j.1755-0238.2012.00188.x

Constable, F. E., Connellan, J., Nicholas, P., Rodoni, B. C. 2013. The reliability of woody indexing for detection of grapevine virus-associated diseases in three different climatic conditions in Australia. Aust. J. Grape Wine Res. 19:74-80. doi:10.1111/j.1755- 0238.2012.00204.x

Coombe, B. G. 1992. Research on development and ripening of the grape berry. Am. J. Enol. Vitic. 43: 101-110.

Daane, K. M., Almeida, R. P. P., Bell, V. A., Botton, M., Fallahzadeh, M., Mani, M., Miano, J. L., Sforza, R., Walton, V. M., and Zaviezo, T. 2012. Biology and management of mealybugs in

21 vineyards. Pages 271-308 in: Arthropod Management in Vineyards. N. J. Bostanian, R. Isaacs, and C. Vincent, eds. Springer, Dordrecht, the Netherlands.

Dawson, W. O., Garnsey, S. M., Tatineni, S., Folimonova, S. Y., Harper, S. J., and Gowda, S. 2013. Front Microbiol. 4.88. doi:10.3389/fmicb.2013.00088. de Saporta, G. 1879. Le monde des plantes avant l’apparition de l’homme. Masson

Dolja, V. V. 2003. Beet yellows virus: the importance of being different. Mol. Plant. Pathol. 4:91-98. doi:10.1046/j.1364-3703.2003.00154.x

Dolja, V. V., Kreuze, J. F., and Valkonen, J. P. T. 2006. Comparative and functional genomics of Closteroviruses. Virus Res. 117:38–51. doi:10.1016/j.virusres.2006.02.002.

Dolja, V. V., and Koonin, E. V. 2013. The closterovirus-derived gene expression and RNA interference vectors as tools for research and plant biotechnology. Front. Microbiol. 4:83. doi:10.3389/fmicb.2013.00083.

Engel, E. A., Escobar, P. F., Rojas,L. A., Rivera, P. A., Fiore, N., and Valenzuela, P. D. 2010. A diagnostic oligonucleotide microarray for simultaneous detection of grapevine viruses. J. Virol. Methods 163:445-451.

Engelbrecht, D. J., and Kasdorf, G. G. F. 1985. Association of a Closterovirus with grapevines indexing positive for grapevine leafroll disease and evidence for its natural spread in grapevine. Phytopathol. Mediterr. 24:101–105.

Engelbrecht, D. J., and Kasdorf, G. G. F. 1990. Transmission of grapevine leafroll disease and associated closteroviruses by the vine mealybug Planococcus ficus. Phytophylactica 22:341–346.

FAOSTAT. 2013. Food and agriculture organization of the United Nations Statistics Division. http:// http://faostat3.fao.org/download/Q/QC/E. (Accessed on September 1, 2015).

Fazeli, C. F., and Rezaian, A. M. 2000. Nucleotide sequence and organization of ten open reading frames in the genome of grapevine leafroll-associated virus 1 and identification of three subgenomic RNAs. J. Gen. Virol. 81:605–615. doi:10.1099/0022-1317-81-3-605.

Fei, F., Lyu, M-D., Li, J., Fan, Z-F., and Cheng, Y-Q. 2013. Complete nucleotide sequence of a Chinese isolate of reveals a 5’UTR of 802 nucleotides. Virus genes 46:182-185. doi:10.1007/ s11262-012-0823-9.

Forsline, P. L., Hoch, J., Limboy, W. F., McFerson, J. R., Golino, D., and Gonsalves, D. 1996. Comparative effectiveness of symptomatology and ELISA for detecting two isolates of grapevine leafroll in Cabernet Franc. Am. J. Enol. Vitic. 47:239-243.

Gale, G. 2002. Saving the vine from phylloxera: a never ending battle. In Wine: A Scientific Exploration, eds. J. Sandler and R. Pidler (London: Taylor and Francis), pp. 70–91.

22

Ghanem-Sabanadzovic, N. A., Sabanadzovic, S., Uyemoto, J. K., Golino, D., and Rowhani, A. 2010. A putative new Ampelovirus associated with grapevine leafroll disease. Arch. Virol. 155:1871–1876. doi:10.1007/s00705-010-0773-2.

Ghanem-Sabanadzovic, N. A., Sabanadzovic, S., Gugerli, P., and Rowhani, A. 2012. Genome organization, serology and phylogeny of grapevine leafroll-associated viruses 4 and 6: Taxonomic implications. Virus Res. 163:120–128. doi:10.1016/j.virusres.2011.09.001.

Giampetruzzi, A. V., Roumi, R., Roberto, U., Malossini, N., Yoshikawa, P., La Notte, P., Terlizzi, F., Credi, R., and Saldarelli, P. 2012. A new grapevine virus discovered by deep sequencing of virus- and viroid derived small RNAs in cv Pinot gris. Virus Res. 163:262-268.

Goheen, A. C. 1970. Grapevine Leafroll. In: Frazier, N.W., Ed. Virus Diseases of Small Fruits and Grapevines. Division of Agricultural Sciences, University of California. pp. 209–212.

Golino, D. A., Sim, S., and Rowhani, A. 2002. Grapevine leafroll disease can be spread by California mealybugs. California Agriculture 56:196-201.

Golino, D. A., Weber, E. A., Sim, S. T., and Rowhani, A. 2008. Leafroll disease is spreading rapidly in a Napa Valley vineyard. Calif. Agric. 62:156–160.

Golino, D. A., Wolpert, J., Sim, S. T., Benz, J., Anderson, M., and Rowhani, A. 2009a. Virus effects on vine growth and fruit components of three California ‘Heritage’ clones of Cabernet Sauvignon. Pages 243-244 in: Proc. XVI Int. Counc. Study Viruses Virus-Like Dis. Grapevine. Dijon, France.

Golino, D. A., Wolpert, J., Sim, S. T., Benz, J., Anderson, M., and Rowhani, A. 2009b. Virus effects on vine growth and fruit components of Cabernet Sauvignon on six rootstocks. Pages 245-246 in: Extended Abstr. 16th Meet. ICVG. Dijon, France.

Gorbalenya, A. E., Koonin, E. V., and Wolf, Y. I. 1990. A new superfamily of putative NTP- binding domains encoded by genomes of small DNA and RNA viruses. FEBS Lett. 262:145– 148. doi:10.1016/0014-5793(90)80175-I.

Gorbalenya, A. E., Koonin, E. V., and Lai, M. M.-C. 1991. Putative papain-related thiol proteases of positive-strand RNA viruses. Identification of rubi- and aphthovirus proteases and delineation of a novel conserved domain associated with proteases of rubi-, α- and coronaviruses. FEBS Lett. 288:201–205. doi:10.1016/0014-5793(91)81034-6.

Goregaoker, S. P., and Culver, J. N. 2003. Oligomerization and activity of the helicase domain of the Tobacco mosaic virus 126- and 183-kilodalton replicase proteins. J. Virol. 77: 3549–3556. doi:10.1128/JVI.77.6.3549-3556.2003.

23 Goszczynski, D. E., Kasdorf, G. G. F., Pietersen, G., and Van Tonder, H. 1996. Grapevine leafroll-associated virus-2 (GLRaV-2) - mechanical transmission, purification, production and properties of antisera, detection by ELISA. S Afr. J. Enol. Vitic. 17:15-26.

Gouveia, P., Dandlen, S., Costa, Â., Marques, N., and Nolasco, G. 2012. Identification of an RNA silencing suppressor encoded by grapevine leafroll-associated virus 3. Eur. J. Plant Pathol. 133:237–245. doi:10.1007/s10658-011-9876-1.

Gribaudo, I., Gambino, G., Bertin, S., Bosco, D., Cotroneo, A., and Mannini, F. 2009. Monitoring the spread of viruses after vineyard replanting with heat-treated clones of Vitis Vinifera “Nebbiolo. J. Plant Pathol. 91:741–744. doi:10.4454/jpp.v91i3.572.

Gugerli, P., Brugger, J. J., and Bovey, R. 1984. L’enroulement de la vigne: mise en évidence de particules virales et développement d’une méthode immunoenzymatique pour le diagnostic rapide. Rev. Suisse Vitic. Arboric Hortic. 16:299–304.

Gugerli, P., and Ramel, M. E. 1993. “Grapevine leafroll-associated virus II analysed by monoclonal antibodies,” in Proceedings of the 11th Congress of ICVG 5–10 September (Montreux: Federal Agricultural Research Station of Changins), 23–24.

Gugerli, P. 2003. Grapevine leafroll and related viruses. Pages 25-31 in: Proc. XIV Int. Counc. Study Viruses Virus-Like Dis. Grapevine. Locorotondo (Bari), Italy.

Gushchin, V. A., Solovyev, A. G., Erokhina, T. N., Morozov, S. Y., and Agranovsky, A. A. 2013. Beet yellows virus replicase and replicative compartments: parallels with other RNA viruses. Front Microbiol. 4:38. doi:10.3389/fmicb.2013.00038.

Gutha, L. R., Casassa, L. F., Harbertson, J. F., and Naidu, R. A. 2010. Modulation of flavonoid biosynthetic pathway genes and anthcyanins due to virus infection in grapevine (Vitis vinifera L.) leaves. BMC Plant Biol. 10:187.

Habili, N., Fazeli, C. F., Ewart, A., Hamilton, R., Cirami, R., Saldarelli, P., Minafra, A., and Rezaian, M. A. 1995. Natural spread and molecular analysis of grapevine leafroll-associated virus 3 in Australia. Phytopathology 85, 1418–1422.

Habili, N., and Nutter, F. W. 1997. Temporal and Spatial Analysis of grapevine leafroll- associated virus 3 in Pinot Noir Grapevines in Australia. Plant Dis. 81:625–628. doi:10.1094/PDIS.1997.81.6.625.

Habili, N., Randles, J. W., and Rowhani, A. 2003. Evidence for the apparent spread of Grapevine virus A and Grapevine leafroll associated virus 9 in a research vineyard in Australia. Extended abstracts 14th Meeting of ICVG, Locorotondo, Italy. 213-214.

Habili, N., Kominek, P., and Little, A. 2007. Grapevine leafroll-associated virus 1 as a common grapevine pathogen. Plant viruses 1:63-68.

24 He, X.-H., Rao, A. L. N., and Creamer, R. 1997. Characterization of beet yellows closterovirus- specific RNAs in infected plants and protoplasts. Phytopathology 87, 347–352. doi:10.1094/PHYTO.1997.87.3.347.

He, F., Mu, L., Yan, G. L., Liang, N. N., Pan, Q. H, Wang, J., Reeves, M. J., and Duan C-Q. 2010. Biosynthesis of anthocyanins and their regulation in colored grapes. Molecules 15:9057– 9091. doi:10.3390/molecules15129057.

Hewitt, W. B. 1954. Some virus and virus-like diseases of grapevines. Bull. Dept. of Agric., State of California, XLII:47-64.

Hilf, M. E., Karasev, A. V., Pappu, H. R., Gumpf, D. J., Niblett, C. L., and Garnsey, S. M. 1995. Characterization of citrus tristeza virus subgenomic RNAs in infected tissue. Virology 208:576– 582.

Hommay, G., Komar, V., Lemaire, O., and Herrbach, E. 2008. Grapevine virus A transmission by larvae of Parthenolecanium corni. Eur. J. Plant Pathol. 121:185–188. doi:10.1007/s10658- 007-9244-3.

Hu, J. S., Gonsalves, D., and Teliz, D. 1990. Characterization of closterovirus-like particles associated with grapevine leafroll disease. J. Phytopathol. 128:1–14.

Jarugula, S., Alabi, O. J., Martin, R. R., and Naidu, R. A. 2010a. Genetic variability of natural populations of grapevine leafroll-associated virus 2 in Pacific Northwest vineyards. Phytopathol. 100:698–707. doi:10.1094/PHYTO-100-7-0698.

Jarugula, S., Gowda, S., Dawson, W. O., and Naidu, R. A. 2010b. 3’-coterminal subgenomic RNAs and putative cis-acting elements of grapevine leafroll-associated virus 3 reveals “unique” features of gene expression strategy in the genus Ampelovirus. Virol. J. 7:180. doi:10.1186/1743- 422X-7-180.

Jelkmann, W., Mikona, C., Turturo, C., Navarro, B., Rott, M. E., Menzel, W., Saldarelli, P., Minafra, A., and Martelli, G. P. 2012. Molecular characterization and taxonomy of grapevine leafroll-associated virus 7. Arch. Virol. 157:359–362. doi:10.1007/s00705-011-1176-8.

Jooste, A. E. C., Maree, H. J., Bellstedt, D. U., Goszczynski, D. E., Pietersen, G., and Burger, J. T. 2010. Three genetic grapevine leafroll-associated virus 3 variants identified from South African vineyards show high variability in their 5′UTR. Arch. Virol. 155:1997–2006. doi:10.1007/s00705-010-0793-y.

Karasev, A. V., Agranovsky, A. A., Rogov, V. V., Miroshnichenko, N. A., Dolja, V. V., and Atabekov, J. G. 1989. Virion RNA of beet yellows closterovirus: cell-free translation and some properties. J. Gen. Virol. 70:241–245.

Karasev, A. V. 2000. Genetic diversity and evolution of closteroviruses.Annu. Rev. Phytopathol. 38:293–324. doi:10.1146/annurev.phyto.38.1.293.

25

Komar, V., Vigne, E., Demangeat, G., Lemaire, O., and Fuchs, M. 2010. Comparative performance analysis of virus-infected Vitis vinifera cv. Savagnin rose grafted onto three rootstocks. Am. J. Enol. Vitic. 61:68–73.

Kovacs, L. G., Hanami, H., Fortenberry, M., and Kaps, M. L. 2001. Latent infection by leafroll agent GLRaV-3 is linked to lower fruit quality in French-American hybrid grapevines Vidal blanc and St. Vincent. Am. J. Enol. Vitic. 52:254-259.

Kobayashi, S., Goto-Yamamoto, N., and Hirochika, H. 2004. Retrotransposon-induced mutations in grape skin color. Science. 14:304(5673):982.

Krenz, B., Thompson, J. R., Fuchs, M., and Perry, K. L. 2012. Complete genome sequence of a new circular DNA virus from grapevine. J. Virol. 86:7715.

Krüger, K., and Douglas, N. 2013. Grapevine leafroll-associated virus 3 (GLRaV-3) transmission by three soft scale insect species (Hemiptera: Coccidae) with notes on their biology. Afr. Entomol.21, 1–8.

Laucou, V., Lacombe, T., Dechesne, F., Siret, R., Bruno, J-P., Dessup, M., Dessup, T., Ortigosa, P., Parra, P., Roux, C., Santoni, S., Vares, D., Peros, J-P., Boursiquot, J-M., and This, P. 2011. High throughput analysis of grape genetic diversity as a tool for germplasm collection management. Theor. Appl. Genet. 122:1233–1245. doi:10.1007/s00122-010-1527-y.

Lee, J., and Martin, R. R. 2009. Influence of grapevine leafroll associated viruses (GLRaV-2 and -3) on the fruit composition of Oregon Vitis vinifera L. cv. Pinot noir: phenolics. Food Chem.112:889–96. doi:10.1016/j.foodchem.2008.06.065.

Lee, J., Keller, K. E., Rennaker, C., and Martin, R. R. 2009. Influence of grapevine leafroll associated viruses (GLRaV-2 and -3) on the fruit composition of Oregon Vitis vinifera L. cv. Pinot noir: free amino acids, sugars, and organic acids. Food Chem.117:99–105. doi:10.1016/j.foodchem.2008.06.065.

Ling, K. S., Zhu, H. Y., Petrovic, N., and Gonsalves, D. 2001. Comparative effectiveness of ELISA and RT-PCR for detecting Grapevine leafroll-associated closterovirus-3 in field samples. Am. J. Enol. Vitic. 52:21-27.

Ling, K. S., Zhu, H. Y., and Gonsalves, D. 2004. Complete nucleotide sequence and genome organization of grapevine leafroll-associated virus 3, type member of the genus Ampelovirus. J. Gen. Virol. 85:2099–2102. doi: 10.1099/vir.0.80007-0.

Liu, Y. P., Peremyslov, V. V., Medina, V., and Dolja, V. V. 2009. Tandem leader proteases of grapevine leafroll-associated virus 2: Host-specific functions in the infection cycle. Virology 383:291–299. doi:10.1016/j.virol.2008.09.035.

26 Lu, R., Folimonov, A., Shintaku, M., Li, W.-X., Falk, B. W., Dawson, W. O., and Ding, S-W. 2004. Three distinct suppressors of RNA silencing encoded by a 20-kb viral RNA genome. Proc. Natl. Acad. Sci. U. S. A. 101:15742–15747. doi:10.1073/pnas.0404940101.

Le Maguet, J., Beuve, M., Herrbach, E., and Lemaire, O. 2012. Transmission of six Ampeloviruses and two vitiviruses to grapevine by Phenacoccus aceris. Phytopathology. 102:717–23. doi:10.1094/PHYTO-10-11-0289.

Le Maguet, J., Fuchs, J. J., Beuve, M., Chadoeuf, J., Herrbach, E., and Lemaire, O. 2013. The role of the mealybug Phenacoccus aceris in the epidemic of Grapevine leafroll associated virus- 1 (GLRaV-1) in two French vineyards. Eur. J. Plant Pathol. 135:415–427. doi:10.1007/s10658- 012-0099-x.

López-Fabuela I, Wetzelb T, Bertolinia E, Basslerb A, Vidala E, Torresa LB, Yustec A, Olmos A. 2013. Real-time multiplex RT-PCR for the simultaneous detection of the five main grapevine viruses. J. Virol. Methods. 188(1-2): 21-24.

Mahfoudhi, N., Digiaro, M., and Dhouibi, M. H. 2009. Transmission of grapevine leafroll viruses by Planococcus ficus (Hemiptera: Pseudococcidae) and Ceroplastes rusci (Hemiptera: Coccidae). Plant Dis. 93:999–1002. doi:10.1094/PDIS-93-10-0999.

Maliogka, V. I., Dovas, C. I., and Katis, N. I. 2008. Evolutionary relationships of virus species belonging to a distinct lineage within the Ampelovirus genus. Virus Res 135:125–135. doi:10.1016/j.virusres.2008.02.015.

Maliogka, V. I., Dovas, C. I., Lotos, L., Efthimiou, K., and Katis, N. I. 2009. Complete genome analysis and immunodetection of a member of a novel virus species belonging to the genus Ampelovirus. Arch. Virol. 154:209–218. doi:10.1007/s00705-008-0290-8.

Maree, H. J., Freeborough, M-J., Burger, J. T. 2008. Complete nucleotide sequence of a South African isolate of grapevine leafroll-associated virus 3 reveals a 5’UTR of 737 nucleotides. Arch. Virol. 153:755-657. doi:10.1007/s00705-008-0040-y.

Maree, H. J., Almeida, R. P. P., Bester, R., Chooi, K. M., Cohen, D., Dolja, V. V., Fuchs, M. F., Golino, D. A., Jooste, A. E. C., Martelli, G. P., Naidu, R. A., Rowhani, A., Saldarelli, P., and Burger, J. T. 2013. Grapevine leafroll-associated virus 3. Front. Microbiol. 82:1-21. doi:10.3389/fmicb.2013.00082.

Martelli, G. P. 1993. Graft-transmissible diseases of grapevines: Handbook for detection and diagnosis. Food and Agriculture Organization of the United Nations in cooperation with the International Council for the Study of Viruses and Virus-Like Diseases of Grapevine. Rome.

Martelli, G. P. 2000. Major graft-transmissible diseases of grapevines: nature, diagnosis, and sanitation. Pages 231-236 in: Proc. 50th Anniv. Annu. Meeting ASEV, Seattle, Washington, USA.

27 Martelli, G. P., Agranovsky, A. A., Bar-Joseph, M., Boscia, D., Candresse, T., Coutts, R. H., Dolja, V. V., Falk, B. W., Gonsalves, D., Jelkmann, W., Karasev, A. V., Minafra, A., Namba, S., Vetten, H. J., Wisler, G. C., and Yoshikawa, N. 2002. The family Closteroviridae revised. Arch. Virol. 147:2039–2044. doi:10.1007/s007050200048.

Martelli, G. P., and Boudon-Padieu, E. 2006. Directory of infectious diseases of grapevines and viruses and virus-like diseases of the grapevine: Bibliographic report 1998–2004, Options Me´diterrane´ennes B55:11-201.

Martelli, G. P., Adams, M. J., Kreuze, J. F., and Dolja, V. V. 2007. Family Flexiviridae: A case study in virion and genome plasticity. Annu. Rev. Phytopathol. 45:73-100. Doi:10.1146/annurev.phyto.45.062806.094401.

Martelli, G. P., Agranovsky, A. A., Bar-Joseph, M., Boscia, D., Candresse, T., Coutts, R. H., Dolja V. V., Hu J. S., Jelkmann W., Karasev A. V., Martin R. R., Minafra A., Namba S., and Vetten H. J. 2011. Family Closteroviridae. In: King A., Adams M. J., Carstens E. B., Lefkowitz E. (eds). Virus Taxonomy. Ninth Report of the International Committee on Taxonomy of Viruses, pp. 987-1001. Elsevier-Academic Press, Amsterdam, the Netherlands.

Martelli, G. P. 2012. Grape virology highlights: 2010-2012. Pages 13-31 in: Proc. 17th Congr. Int. Counc. Study Virus Virus-like Dis. Grapevine (ICVG), Davis, California, USA.

Martelli, G. P., Ghanem-sabanadzovic, N. A., Agranovsky, A. A., Rwahnih, M. A, Dolja, V. V., and Dovas, C. I. 2012. Taxonomic revision of the family Closteroviridae with special reference to the grapevine leafroll-associated members of the genus Ampelovirus and the putative species unassigned to the family. J. Plant Pathol. 94:7–19. doi:10.4454/jpp.fa.2012.022.

Martelli, G. P. 2014. Directory of virus and virus-like diseases of the grapevine and their agents. J. Plant Pathol. 96(1S):1–136. doi:10.4454/JPP.V96I1SUP.

Martin, R. R., Eastwell, K. C., Wagner, A., Lamprecht, S., and Tzanetakis, I. E. 2005. Survey for viruses of grapevine in Oregon and Washington. Plant Dis. 89:763–766. doi:10.1094/PD-89- 0763.

McGovern, P.E. 2004. Ancient wine: the search for the origins of viniculture. Princeton University Press.

Mekuria, T. A., Gutha, L. R., Martin, R. R., and Naidu, R. A. 2009. Genome diversity and intra and interspecies recombination events in Grapevine fanleaf virus. Phytopath. 99:1394-1402.

Mikona, C., and Jelkmann, W. 2010. Replication of grapevine leafroll-associated virus 7 (GLRaV-7) by Cuscuta species and its transmission to herbaceous plants. Plant Dis. 94:471–476. doi:10.1094/PDIS-94-4-0471.

Mullins, M. G., Bouquet, A., and Williams, L. E. 1992. Biology of the grapevine. Press Syndicate of University of Cambridge. Cambridge, pp. 239.

28

Naidu, R. A. 2011. Virus Update: The Status of Washington Vineyards. Viticulture and enology extension news-Fall 2011. WSU. http://wine.wsu.edu/research-extension/publications/newsletter/ (Accessed on September 1, 2015).

Naidu, R. A., Rowhani, A., Fuchs, M., Golino, D., and Martelli, G. P. 2014. Grapevine leafroll: a complex viral disease affecting a high-value fruit crop. Plant Dis.98:1172–85. doi:10.1094/PDIS- 08-13-0880-FE.

Naidu, R. A., and Walsh, D. Is ‘grape virus tax’ hitting your pocketbook? Good Fruit Grower. May 15, 2015. Vol. 66, No. 10, pp 10-11.

Naidu, R. A., Maree, H. J., and Burger, J. T. 2015. Grapevine leafroll disease and associated viruses: A unique pathosystem. Annu. Rev. Phytopathol. 53:613-34. doi:10.1146/annurev- phyto-102313-045946.

Namba, S., Yamashita, S., Doi, Y., Yora, K., Terai, Y., and Yano, R. 1979. Grapevine leafroll virus, a possible member of closteroviruses. Ann. Phytopathol. Soc. Jpn. 45:497–502. doi:10.3186/jjphytopath.45.497.

Napuli, A. J., Alzhanova, D. V., Doneanu, C. E., Barofsky, D. F., Koonin, E. V, and Dolja, V. V. 2003. The 64-kilodalton capsid protein homolog of Beet yellows virus is required for assembly of virion tails. J. Virol. 77:2377–2384. doi:10.1128/JVI.77.4.2377-2384.2003.

NASS. 2015. National Agricultural Statistics Service. http://www.nass.usda.gov/Publications (Accessed on September 1, 2015).

OIV. 2013. International Organization of Vine and Wine: Statistical Report on World Vitiviniculture, pp. 28.

Oliver, J. E., and Fuchs, M. 2011. Tolerance and resistance to viruses and their vectors in Vitis sp.: A virologist’s perspective of the literature. Am. J. Enol. Vitic. 62:438–451. doi:10.5344/ajev.2011.11036.

Osman, F., Leutenegger, C., Golino, D., and Rowhani, A. 2007. Real-time RT-PCR (TaqMan ®) assays for the detection of grapevine leafroll associated viruses 1–5 and 9. J. Virol. Methods 141:22-29.

Osman, F., Olineka, T., Hodzic, E., Golino, D., and Rowhani, A. 2012. Comparative procedures for sample processing and quantitative PCR detection of grapevine viruses. J. Virol. Methods 179:303-310.

Peng, C. W., Peremyslov, V. V., Mushegian, A. R., Dawson, W. O., and Dolja, V. V. 2001. Functional specialization and evolution of leader proteinases in the family Closteroviridae. J. Virol. 75:12153–12160. doi:10.1128/JVI.75.24.12153-12160.2001.

29 Peremyslov, V. V., Hagiwara, Y, and Dolja V. V. 1999. HSP70 homolog functions in cell-to-cell movement of a plant virus. Proc. Natl. Acad. Sci. USA 96:14771–76. doi:10.1073/pnas.96.26.14771.

Peremyslov, V. V., Andreev, I. A., Prokhnevsky, A. I., Duncan, G. H., Taliansky, M. E., and Dolja, V. V. 2004. Complex molecular architecture of beet yellows virus particles. Proc. Natl. Acad. Sci. U.S.A. 101:5030–5035. doi:10.1073/pnas.0400303101.

Pietersen, G. 2006. Spatio-temporal distribution dynamics of grapevine leafroll disease in Western Cape vineyards. Pages 126-127 in: Proc. 15th Int. Counc. Study Viruses Virus-Like Dis. Grapevine. Stellenbosch, South Africa.

Poojari, S., Alabi, O. J., and Naidu, R. A. 2013a. Molecular characterization and impacts of a strain of grapevine leafroll-associated virus 2 causing asymptomatic infection in a wine grape cultivar. Virol. J. 10:324. doi:10.1186/1743-422X-10-324.

Poojari, S., Alabi, O. J., Fofanov, V. Y., and Naidu, R. A. 2013b. A leafhopper-transmissible DNA virus with novel evolutionary lineage in the family Geminiviridae implicated in grapevine redleaf disease by next-generation sequencing. PLoS ONE 8(6):e64194. doi:10.1371/journal.pone.0064194.

Pourrahim, R., Farzadfar, S. H., Golnaraghi, A. R., and Ahoonmanesh, A. 2007. Partial molecular characterization of some grapevine fanleaf virus isolates from North-east of Iran. J. Phytopathol. 155:754–757. doi:10.1111/j.1439-0434.2007.01299.x.

Prokhnevsky, A. I., Peremyslov, V. V, Napuli, A. J., and Dolja, V. V. 2002. Interaction between long-distance transport factor and Hsp70-related movement protein of beet yellows virus. J. Virol. 76:11003–11011. doi:10.1128/JVI.76.21.11003-11011.2002.

Rayapati, N. A., O’Neil, S., and Walsh, D. 2008. Grapevine Leafroll Disease. WSU Extension Bulletin EB2027E, pp. 20 Available at: http://cru.cahe.wsu.edu/CEPublications/eb2027e/eb2027e.pdf

Reed, J. C., Kasschau, K. D., Prokhnevsky, A. I., Gopinath, K., Pogue, G. P., Carrington, J. C., and Dolja, V. V. 2003. Suppressor of RNA silencing encoded by beet yellows virus. Virol. 306:203–209. doi:10.1016/S0042-6822(02)00051-X.

Ricketts, K. D., Gomez, M. J., Atallah, S. S., Fuchs, M. F., Martinson, T. E., Battany, M. C., Bettiga, L J., Cooper, M. L., Verdegaal, P. S., and Smith, R. J. 2015. Reducing the economic impact of grapevine leafroll disease in California: Identifying optimal disease management strategies. Am. J. Enol. Vitic. 66:138-147. doi:10.5344/ajev.2014.14106.

Rosciglione, B., and Gugerli, P. 1986. Maladies de l’enroulement et du bois strié de la vigne: analyse microscopique et sérologique. Rev. Suisse Vitic. Arboric. Hortic. 18:207–211.

30 Rowhani, A. 1992. Use of (ab’) 2 antibody fragment in ELISA for detection of grapevine viruses. Am. J. Enol. Vitic. 43:38-40.

Rowhani, A., Biardi, L., Johnson, R., Saldarelli, P., Zhang, Y. P., Chin, J., and Green, M. 2000. Simplified sample preparation method and one-tube RT-PCR for grapevine viruses. Pages 82-83 in: Proc. XIII Int. Counc. Study Viruses Virus-Like Dis. Grapevine, Adelaide 2000.

Rowhani, A., Uyemoto, J. K., Golino,D. A., and Martelli, G. P. 2005. Pathogen testing and certification of Vitis and Prunus species. Annu. Rev. Phytopathol. 43:261-278. doi:10.1146/annurev.phyto.43.040204.135919.

Rozanov, M. N., Koonin, E. V., and Gorbalenya, A. E. 1992. Conservation of the putative methyltransferase domain: a hallmark of the “Sindbis-like” supergroup of positive-strand RNA viruses. J. Gen. Virol. 73:2129–2134. doi:10.1099/0022-1317-73-8-2129.

Salmaso, M., Faes, G., Segala, C., Stefanini, M., Salakhutdinov, I., Zyprian, E., Toepfer, R., Grando, M. S., and Velasco, R. 2004. Genome diversity and gene haplotypes in the grapevine (Vitis vinifera L.), as revealed by single nucleotide polymorphisms. Mol. Breed. 14:385–395. doi:10.1007/s11032-005-0261-7.

Satyanarayana, T., Gowda, S., Mawassi, M., Albiach-Martí, M. R., Ayllón, M. A., and Robertson, C., Garnsey, S. M., and Dawson, W. O. 2000. Closterovirus encoded HSP70 homolog and p61 in addition to both coat proteins function in efficient virion assembly. Virology. 278:253–265. doi:10.1006/viro.2000.0638.

Satyanarayana, T., Gowda, S., Ayllón, M. A., and Dawson, W. O. 2004. Closterovirus bipolar virion: evidence for initiation of assembly by minor coat protein and its restriction to the genomic RNA 5’ region. Proc. Natl. Acad. Sci. U. S. A. 101:799–804. doi:10.1073/pnas.0307747100.

Scheu, G. 1935. Die Rollkrankheit des Rebenstockes. Der Deutsche Weinbau 14:222–223. Sforza, R., Boudon-Padieu, E., and Greif, C. 2003. New mealybug species vectoring grapevine leafroll-associated viruses -1 and -3 (GLRaV-1 and -3). Eur. J. Plant Pathol. 109:975–981. doi:10.1023/B:EJPP.0000003750.34458.71.

Schon, I., Martens, K., van Dijk, P., Forneck, A., Benjak, A., and Ruhl, E. 2009. Grapevine (Vitis ssp.): Example of clonal reproduction in agricultural important plants. In: Schon, I., Martens, K., and van Dijk, P (eds) Lost Sex, pp. 581. Springer, The Netherlands.

Sforza R, Boudon-Padieu E, Greif C. 2003. New mealybug species vectoring grapevine leafroll- associated viruses-1 and -3 (GLRaV-1 and -3). Eur. J. Plant Pathol. 109:975-981.

Sim, S. T., Rowhani, A., and Golino, D. A. 2003. Experimental transmission of Grapevine leafroll associated virus 5 and 9 by longtailed mealybugs. Extended abstracts 14th Meeting of ICVG, Locorotondo, Italy. 211-212.

31 Sokolsky, T., Cohen, Y., Zahavi, T., Sapir, G., and Sharon, R. 2013. Potential efficiency of grapevine leafroll disease management strategies using simulation and real spatio-temporal disease infection data. Aust. J. Grape Wine Res. 19:431–438. doi:10.1111/ajgw.12037.

Soule, M. J., Eastwell, K. C., and Naidu, R. A. 2006. First report of Grapevine leafroll associated virus-3 in American Vitis spp. Grapevines in Washington State. Plant Disease. 90:1461.

Sudarshana, M. R., Perry, K. L., and Fuchs, M. F. 2015. Grapevine red blotch-associated virus, an emerging threat to the grapevine industry. Phytopathol. 105:1026-1032. doi:10.1094/PHYTO- 12-14-0369-FI.

Tanne, E., Ben-Dov, Y., and Raccah, B. 1989. Transmission of closterovirus-like particles by mealybugs (Pseudococcidae) in Israel. Phytoparasitica 17:64.

This, P., Lacombe, T., and Thomas, M. R. 2006. Historical origins and genetic diversity of wine grapes. Trends Genet. 22:511–519. doi:10.1016/j.tig.2006.07.008.

Thompson, J. R., Fuchs, M., and Perry, K. L. 2012a. Genomic analysis of grapevine leafroll associated virus-5 and related viruses. Virus Res. 163:19–27. doi:10.1016/j.virusres.2011.08.006.

Thompson, J. R., Fuchs, M., Fischer, K. F., and Perry, K. L. 2012b. Macroarray detection of grapevine leafroll-associated viruses. J. Virol. Methods 183:161-169.

Tian, T., Rubio, L., Yeh, H. H., Crawford, B., and Falk, B. W. 1999. Lettuce infectious yellows virus: invitro acquisition analysis using partially purified virions and the whitefly Bemisia tabaci. J. Gen. Virol. 80:1111-1117. doi:10.1099/0022-1317-80-5-1111.

Troggio, M., Vezzulli, S., Pindo, M., Malacarne, G., Fontana, P., Moreira, F. M., Costantini, L., Grando, M. S., Viola, R., and Velasco, R. 2008. Beyond the genome, opportunities for a modern viticulture: A research overview. Am. J. Enol. Vitic. 59:117–127.

Tsai, C.-W., Rowhani, A., Golino, D. A., Daane, K. M., and Almeida, R. P. P. 2010. Mealybug transmission of Grapevine leafroll viruses: an analysis of virus-vector specificity. Phytopath. 100:830–834. doi:10.1094/PHYTO-100-8-0830. van den Born, E., Omelchenko, M. V., Bekkelund, A., Leihne, V., Koonin, E. V., Dolja, V. V., and Falnes, P. O. 2008. Viral AlkB proteins repair DNA damage by oxidative demethylation. Nucleic Acids Res 36: 5451-5461. doi:10.1093/nar/gkn519.

Vecchi-Staraz, M. D., Laucou, V., Bruno, G., Lacombe, T., Gerber, S., Bourse, T., Boselli, M., and This, P. 2009. Low level of pollen-mediated gene flow from cultivated to wild grapevine: Consequences for the evolution of the endangered subspecies Vitis vinifera L. subsp. silvestris. J. Hered. 100:66–75. doi:10.1093/jhered/esn084.

32 Velasco, L., Cretazzo, E., Padilla, C. V., and Janssen, D. 2015. Grapevine leafroll associated virus 4 strain 9: Complete genome and quantitaive analysis of virus-derived small interfering RNA populations. J. Plant Pathol. 97:189-192. doi:10.4454/JPP.V97I1.051.

Walker, A. R., Lee, E., Bogs, J., McDavid, D. A. J., Thomas, M. R., and Robinson, S. P. 2007. White grapes arose through the mutation of two similar and adjacent regulatory genes. Plant J. 49:767-959. doi:10.1111/j.1365-313X.2006.02997.x.

Walsh, H. A., and Pietersen, G. 2013. Rapid detection of Grapevine leafroll-associated virus type 3 using a reverse transcription loop-mediated amplification. J. Virol. Methods 194:308-316.

Zee, F., Gonsalves, D., Goheen, A., Kim, K. S., Pool, R., and Lee, R. F. 1987. Cytopathology of leafroll-diseased grapevines and the purification and serology of associated closterovirus-like particles. Phytopathology 77:1427–1434.

Zhang, Y., Singh, K., Kaur, R., and Qiu, W. 2011. Association of a novel DNA virus with the grapevine vein-clearing and vine decline syndrome. Phytopathology 101:1081-1090.

Zhu, H. Y., Ling, K. S., Goszczynski, D. E., McFerson, J. R., and Gonsalves, D. 1998. Nucleotide sequence and genome organization of grapevine leafroll-associated virus-2 are similar to beet yellows virus, the closterovirus type member. J. Gen. Virol. 79:1289–1298. doi:10.1099/0022-1317-79-5-1289.

Zimmermann, D., Bass, P., Legin, R., and Walter, B. 1990. Characterization and serological detection of four closterovirus-like particles associated with leafroll disease of grapevines. J. Phytopathol. 130:205–218.

Zohary, D. 1996. Domestication of the grapevine Vitis vinifera L. in the Near East. In the origins and ancient history of wine. Edited by McGovern, P., Fleming, S., and Katz, S. New York: Gordon and Breach, pp. 23–30.

33 CHAPTER TWO

SEQUENCE ANALYSIS OF GRAPEVINE LEAFROLL-ASSOCIATED VIRUS 1

ISOLATES FROM WASHINGTON VINEYARDS

Bhanu Priya Donda, Sridhar Jarugula and Rayapati A. Naidu

(Unpublished)

ABSTRACT

Grapevine leafroll disease (GLD) is one of the most economically important virus disease affecting wine grapes (Vitis vinifera L.) in almost all grapevine-growing regions around the world. Five distinct species of Grapevine leafroll-associated viruses (GLRaVs, family:

Closteroviridae), designated as GLRaV-1, -2, -3, -4, and -7, have been documented in grapevines. Except for GLRaV-1 (genus: Ampelovirus), the complete genome sequence of other

GLRaV species is available. In this study, the complete genome sequence of two GLRaV-1 isolates was determined to be 18,731 (isolate WA-CH) and 18,946 (isolate WA-PN) nucleotides.

The genome of WA-CH and WA-PN contains nine putative open reading frames (ORFs) and their organization was similar to GLRaV-1 sequences reported previously from Australia and

Canada. Although the size of these ORFs was similar between the two isolates, the 5’ and 3’ non-translated regions (NTRs) were distinctly different in size and nucleotide sequence composition. In Northern blots, three of the eight 3’co-terminal subgenomic (sg) RNAs were detected at higher levels and were putatively designated as sgRNAs specific to CP, p21 and p24.

Among them, the sgRNA corresponding to p24 gene accumulated at the highest level, followed by sgRNAs for CP and p21, respectively. The 5’ termini of five putative sgRNAs (viz. CP,

CPd1, CPd2, p21 and p24) were mapped and their leader sequences determined to be 68 (WA-

34 CH)/67 (WAPN), 27, 15, 49 and 18 nt, respectively. Using the 5’NTR sequences of WA-CH and

WA-PN isolates, a reverse transcription polymerase chain reaction and restriction fragment length polymorphism assay was developed to distinguish GLRaV-1 variants in vineyards.

INTRODUCTION

Among the grapevine virus diseases documented so far, grapevine leafroll disease (GLD) is considered the most economically important virus disease of wine grapes (Vitis vinifera) across the many grapevine-growing regions in the world (Martelli, 2014; Naidu et al., 2014).

GLD is known to affect vine vigor, fruit yield, and berry and wine quality attributes (Golino et al., 2009 a & b; Lee and Martin, 2009; Lee et al., 2009; Basso et al., 2010; Komar et al., 2010;

Alabi et al., 2012). One of the unique features of GLD is cultivar-specific differences in symptoms, with red-berried cultivars showing interveinal reddening or reddish purple discoloration and white-berried cultivars showing mild yellowing (Rayapati et al., 2008). These symptoms usually begin to appear after véraison and become apparent as the season advances, with symptomatic leaves of both red- and white-berried cultivars showing downward rolling of leaf margins towards the end of the season.

Several monopartite viruses belonging to the family Closteroviridae, designated serially as Grapevine leafroll-associated virus (GLRaV) 1, 2, 3, etc., have been found in grapevines showing GLD symptoms or suspected for infection with GLD (Martelli et al., 2002; Martelli,

2014; Naidu et al., 2014). Recent taxonomic considerations have classified them into five distinct species; namely, GLRaV-1, -2, -3, -4, and -7, with other GLRaVs grouped as strains of GLRaV-

4 (Martelli et al., 2012). Among them, GLRaV-1, -3, -4 and its strains were assigned to the genus

Ampelovirus, GLRaV-2 to the genus Closterovirus, and GLRaV-7 to the genus Velarivirus (Al

35 Rwahnih et al., 2012; Martelli et al., 2012; Martelli, 2014; Naidu et al., 2015). Thus, the majority of grapevine-infecting closterovirids belong to the genus Ampelovirus. The complete genome sequences of many of these GLRaVs, with the exception of GLRaV-1, have been completed

(Zhu et al., 1998; Ling et al., 2004; Maree et al., 2008; Maliogka et al., 2009; Jarugula et al.,

2010; Jooste et al., 2010; Ghanem-Sabanadzovic et al., 2010, 2012; Bester et al., 2012, 2014;

Jelkmann et al., 2012; Thompson et al., 2012; Fei et al., 2013; Velasco et al., 2015). The available data indicates that the genome size of GLRaV species is variable between 13,626 nucleotides (nt) (viz. GLRaV-4 strain GLRaV-Car, Ghanem-Sabanadzovic et al., 2010) and

18,671 nt (viz. GLRaV-3, Bester et al., 2012). Based on their genome size, grapevine-infecting ampeloviruses were divided into two subgroups: GLRaV-1 and GLRaV-3 having large genome size assigned to subgroup I and GLRaV-4 and its strains with small genomes, relative to

GLRaV-1 and -3, to subgroup II (Martelli et al., 2012). Although many GLRaVs and their strains have been implicated in the induction of GLD symptoms, some strains of GLRaV-2 (Bertazzon et al., 2010; Alkowni et al., 2011; Poojari et al., 2013) and GLRaV-7 (Al Rwahnih et al., 2012;

Jelkmann et al., 2012) causing asymptomatic infections (with no apparent leafrolling symptoms) have been reported. Although all GLRaVs can be disseminated via vegetative propagation and grafting, ampeloviruses and their strains are reported to be transmitted by different species of mealybugs (Hemiptera: Pseudococcidae) and scale insects (Hemiptera: Coccidae), whereas vectors for GLRaV-2 and GLRaV-7 are yet to be known (Almeida et al., 2013; Naidu et al.,

2014 and cited references).

GLRaV-1 is considered as the second most widely distributed virus, after GLRaV-3, across the many grapevine-growing regions (Habili et al., 2007). Unlike other GLRaVs, only partial genome sequence of GLRaV-1 is available from Australia (Fazeli and Rezaian, 2000;

36 AF195822) and Canada (NC_016509). Based on the available genome sequence, GLRaV-1 encodes nine open reading frames (Fig. 4A; Fazeli and Rezaian, 2000). The ORF 1 encodes two replication-associated proteins that constitutes the ‘replication gene block’ (RGB), analogous to other monopartite viruses in the family Closteroviridae (Dolja et al., 2006). The remaining eight

ORFs are located downstream of the RGB towards the 3’ terminus of the virus genome. The first five of these eight ORFs, designated consecutively from 5’ to 3’ end of the virus genome, are p7, heat-shock protein 70 homolog (HSP70h), p55, coat protein (CP) and the first divergent copy of the CP (CPd1). These ORFs comprise the ‘quintuple gene block’ (QGB), similar to other monopartite closteroviruses (Dolja et al., 2006; Naidu et al., 2015). Downstream of the QGB are three ORFs, designated sequentially as CPd2, p21 and p24 that are unique to GLRaV-1. Unlike other monopartite closteroviruses, including GLRaV-2, -3, and -7, GLRaV-1 encodes two divergent copies of the CP, CPd1 and CPd2. This unique feature sets GLRaV-1 apart from other closteroviruses.

Relative to GLRaV-3, limited progress has been made in understanding the molecular biology of GLRaV-1 (Fazeli and Rezaian, 2000; Habili et al., 2007). A few studies have shown that GLRaV-1 can occur as genetically distinct variants, with CPd2 as the most divergent compared to other genes, such as CP and p24 (Little et al., 2001; Alabi et al., 2011). Since full- genome sequence of a virus is essential for addressing fundamental questions regarding molecular biology, host-virus interactions and epidemiology, this study was undertaken to determine the complete genome sequence of GLRaV-1 from Washington vineyards. The results indicated that, although the overall genome organization is similar to previous reports (Fazeli and

Rezaian, 2000; Little, 2004), the most striking novel feature of GLRaV-1 isolates determined in this study are their large genome size and an unusually long and variable non-translated region

37 (NTR) at the 5’-terminus of the genome, compared to other characterized genomes of closteroviruses. In addition, the 5’-termini of subgenomic (sg) RNAs specific to CP, CPd1,

CPd2, p21, and p24 were mapped in the GLRaV-1 genome and their leader sequences determined to examine comparative transcriptional strategy of the virus relative to other monopartite closteroviruses. A portion of this work was presented at the Annual Meeting of the

American Phytopathological Society at Pasadena, CA, during August 1-5, 2015 (Donda and

Naidu, 2015).

MATERIALS AND METHODS

Source materials and isolation of viral double-stranded RNA

GLRaV-1 isolates used in this study were obtained from two wine grape cultivars (V. vinifera, cvs. Chardonnay and Pinot Noir) planted in two geographically separate commercial vineyard blocks and two ornamental grapevine cultivars (V. vinifera, cv. Purpurea and V. californicum, cv. Roger’s Red) maintained in the greenhouse (Alabi et al., 2011). Cambial scrapings of mature canes from cvs. Chardonnay and Pinot Noir were used for isolating genomic-length replicative-form of double-stranded (ds) RNA, essentially as described by

Valverde et al. (1990) and Jarugula et al. (2010). The integrity of dsRNA preparations was verified by resolving in 0.8% agarose gels pre-stained with GelRed and visualizing under UV light using a transilluminator (Bio-Rad Universal Hood, Bio-Rad Laboratories). The dsRNA preparations were tested by RT-PCR to confirm the presence of GLRaV-1 using primers specific to the HSP70h and CP as described earlier (Alabi et al., 2011).

cDNA synthesis, RT-PCR, cloning and sequencing

38 The dsRNA preparations were used as a template for amplification of the genome of

GLRaV-1, similar to the amplicons-based strategy reported for cloning the genome of GLRaV-3

(Jarugula et al., 2010). Initially, consensus primers were designed based on GLRaV-1 sequences available in GenBank (NC_016509 and AF195822). Subsequently, primers were designed based on the sequence of GLRaV-1 isolates (designated as WA-CH and WA-PN) generated in this study. Primer sequences and their location in the genome of GLRaV-1 isolate WA-CH are listed in the Table 1. The denatured dsRNA preparation was used as a template for cDNA synthesis using primers complementary to specific regions of the virus genome. The assay was carried out in 20 µL reaction mixture in the presence of Superscript III reverse transcriptase (ThermoFischer

Scientific, Grand Island, New York), according to the manufacturer’s instructions. The cDNA preparation served as a template in subsequent PCR assays for amplifying different portions of the virus genome. Concentration of the cDNA was measured using NanoDropTM 2000c spectrophotometer (ThermoFischer Scientific, Grand Island, New York). Specific primer pairs were used to generate overlapping DNA fragments covering most of the virus genome (Fig. 4B).

Each PCR reaction performed in a 50μL reaction mixture consisted of a final concentration of 2 to 5 ng of cDNA template, 200μM each dNTPs, 0.3μM each of virus specific primers, 1x concentration of PrimeSTAR GXL reaction buffer with 1mM Mg+2 and 1.25U of PrimeSTAR

GXL DNA polymerase (Takara Bio Inc., Japan). PCR conditions included an initial denaturation at 94°C for 30 sec, followed by 35 cycles of denaturation at 98°C for 10 sec, primer annealing at

60°C for 5 sec, primer extension at 72°C for 1 min per 1kb PCR product, and 1 cycle of final extension at 72°C for 10 min. An aliquot of each PCR assay product was resolved in 0.8% agarose gels and DNA bands were visualized under UV light using a transilluminator mentioned above. The amplicons were gel purified with GeneClean® III Kit (MP Biomedicals, Santa Ana,

39 California), mixed with dA-tailing reaction mixture containing 10mM Tris-HCl, 1.5mM MgCl2,

50mM KCl, pH 8.3, 0.2mM dATP, 1U of Taq DNA polymerase (Roche, Mannheim, Germany) and incubated at 72°C for 20 min. The dA-tailed amplicons were subsequently cloned into pCR®-XL-TOPO® vector (ThermoFischer Scientific, Grand Island, New York) and transformed into chemically competent Escherichia coli cells (ThermoFischer Scientific, Grand Island, New

York), according to the manufacturer’s instructions. Colony PCR was used to identify E. coli colonies harboring recombinant DNA clones using blue-white screening method and the principle of α-complementation (Ullmann et al., 1967; Langley et al., 1975; Vieira and Messing,

1982). Recombinant plasmid DNA obtained from three independent colonies per amplicon was sequenced in both orientations at Retrogen, Inc. (San Diego, California). Additional clones were sequenced as needed to resolve sequence ambiguities between clones representing each amplicon.

Figure 4. Schematic representation of the strategy used for cloning the complete genome of

GLRaV-1. (A) Genome organization of GLRaV-1. The ORFs are shown as boxes acros the genome and labeled with either associated protein designations or with approximate molecular

40 weight and a common “p” designator (Dolja et al., 2006). L-Pro, papain-like leader proteinase;

MET, methyl transferase domain; AlkB, an AlkB domain; HEL, RNA helicase domain; RdRp,

RNA-dependent RNA polymerase; p7, a 7-kDa protein; HSP70h, heat shock protein-70 homologue; p55, 55-kDa protein; CP, coat protein; CPd1 and CPd2, divergent copies of the CP, respectively; p21, 21-kDa protein; p24, 24-kDa protein. (B) The location of primers used for RT-

PCR amplification of specific regions of the genome and size of amplicons. The nucleotide sequences of primers VA-F, VA-R, VB-F, VB-R, VC-F, VC-R, VD-F and VD-R are listed in

Table 1.

Determination of 5’ and 3’ terminal sequences

The 5’ terminal sequence of GLRaV-1 was determined by rapid amplification of cDNA ends using a commercial 5’RACE system (Version 2.0; ThermoFischer Scientific, Grand Island,

New York), according to the manufacturer’s instructions. Total RNA was extracted from cambial scrapings of grapevines tested positive for GLRaV-1 using Sigma Spectrum™ Plant Total RNA

Kit (Sigma-Aldrich, St. Louis, Missouri), according to the manufacturer’s instructions. First- strand cDNA was synthesized from total RNA using the primer 5’

CAAGAAGACTTGATTTCCAT 3’, complementary to 882 nt to 901 nt of GLRaV-1 isolate

WA-CH (Table 2). A homopolymeric dC tail was ligated to the 3’ termini of purified cDNA using terminal deoxynucleotidyl transferase (ThermoFischer Scientific, Grand Island, New

York). The dC-tailed cDNA was subsequently used in PCR to amplify virus-specific DNA fragments using two sets of primer pairs. The first primer pair consisted of a virus-specific primer 5’ GCGTGCTTCACTTTATAGTATCCCTCCTC 3’, complementary to 649 nt to 677 nt in GLRaV-1 isolate WA-CH, and the abridged anchor primer (AAP) supplied with the 5’RACE

41 kit (Version 2.0; ThermoFischer Scientific, Grand Island, New York). The second pair consisted of virus-specific primer 5’ TCGTTTCCTTTCACTCAGGGACGA 3’, complementary to 372 nt to 395 nt in GLRaV-1 isolate WA-CH, and the abridged universal amplification primer (AUAP) supplied with the 5’RACE kit (Version 2.0; ThermoFischer Scientific, Grand Island, New York).

The PCR conditions included an initial denaturation at 94°C for 3 min, followed by 35 cycles of denaturation at 94°C for 10 sec, primer annealing at 56°C for 30 sec and extension at 72°C for 1 min, and a final extension step at 72°C for 5 min. For additional verification of 5’end sequence of GLRaV-1, cDNA obtained from dsRNA using SuperScriptIII reverse transcriptase

(Invitrogen, Carlsbad, California) and 5’ CAAGAAGACTTGATTTCCAT 3’, complementary to

882 nt to 901 nt of GLRaV-1 isolate WA-CH (Table 2) was dA-tailed with Yeast Poly(A) polymerase (USB, Cleveland, Ohio). The dA-tailed cDNA was subsequently used in PCR to amplify virus-specific DNA using two primer pairs. The first primer pair consisted of a modified

AAP primer with oligo dT sequence 5’

GGCCACGCGTCGACTAGTACTTTTTTTTTTTTTTTT 3’ and a virus-specific inner primer 5’

GCGTGCTTCACTTTATAGTATCCCTCCTC 3’ complementary to 649 nt to 677 nt in

GLRaV-1 isolate WA-CH. The second pair consisted of virus-specific inner primer 5’

TCGTTTCCTTTCACTCAGGGACGA 3’, complementary to 372 nt to 395 nt in GLRaV-1 isolate WA-CH, and the abridged universal amplification primer (AUAP) supplied with the

5’RACE kit (Version 2.0; ThermoFischer Scientific, Grand Island, New York). The PCR conditions for this reaction is same as described above for determining 5’ end using dC-tailed cDNA.

For determining 3’end sequence, dA tailing of the 3’-end was employed using Yeast

Poly(A) polymerase (USB, Cleveland, Ohio) as described above. Additionally, 5’RACE method

42 was performed using dsRNA preparations and virus-specific outer and inner primers designed to amplify the minus strand of dsRNA (Table 2). PCR reaction was performed according to the protocol described in the 5’RACE kit (ThermoFischer Scientific, Grand Island, New York). An aliquot of each PCR assay product was resolved in 1.5 % agarose gels pre-stained with GelRed and visualized under a UV light using a transilluminator as described above. The resulting amplicons were cloned into pCR®2.1-TOPO® vector (ThermoFischer Scientific, Grand Island,

New York) as described above. Fifteen independent clones per amplicon were sequenced in both the orientations at Retrogen, Inc. (San Diego, California).

RT-PCR based RFLP method for molecular typing of GLRaV-1 isolates

The 5’NTR sequences of GLRaV-1 isolates obtained in this study were screened in silico for unique restriction enzyme sites using NEBcutter V2.0 (New England Biolabs, Ipswich,

Massachusetts). Amplicons of approximately 900 nt length, covering the 5’NTR and 44 nt of the

N-terminal portion of the ORF1a, were generated by RT-PCR using primers VA-F (5’

GTATTGTCCTAGTAGGTA 3’), corresponding to 1 to 18 nt, and 5’-OP (5’

CAAGAAGACTTGATTTCCAT 3’), complementary to 882 to 901 nt, of GLRaV-1 isolate WA-

CH (Tables 1 and 2) and subjected to digestion with XhoI (New England Biolabs, Ipswich,

Massachusetts). The DNA fragments were resolved in 1.5% agarose gels and visualized under

UV light using a transilluminator as described above.

Northern blot hybridization

Total RNA was isolated from cambial scrapings of canes collected from GLRaV-1- positive Chardonnay vines using Sigma Spectrum™ Plant Total RNA Kit (Sigma-Aldrich, St.

43 Louis, Missouri), according to the manufacturer’s instructions. Total RNA was resolved in formaldehyde denaturing 1.1% agarose gels in MOPS buffer and transferred onto a nylon membrane using an electrotransfer unit (Hoefer Pharmacia Biotech, San Francisco, California).

Prehybridization and subsequent hybridization using nonradioactive digoxigenin (DIG)-labelled riboprobes were carried out essentially as described in Jarugula et al. (2010). For preparing the probe, sequences specific to the p24 ORF was amplified from a cDNA clone using the primer sequence 5’-ATGGCGTCACTTATACCTAAGTATGTC 3’, corresponding to 17,643 to 17,669 nt, and complementary primer 3’-

ATTTAGGTGACACTATAGCACACCAAATTGCTAGCGATAGC 5’, corresponding to 18,274 to 18,252 nt in the genome of GLRaV-1 isolate WA-CH. The complementary primer contained

SP6 polymerase promoter sequence (indicated in Italics). The approximately 630 nt DNA fragment amplified by PCR was gel purified with GeneClean® III Kit, according to the manufacturer’s instructions (MP Biomedicals, Santa Ana, California). The purified DNA fragment was used as a template to generate nonradioactive DIG-labeled riboprobes using SP6-

RNA polymerase and nucleotides containing DIG-labeled UTP (DIG RNA labeling kit (SP6/T7),

Sigma-Aldrich, St. Louis, Missouri), according to the manufacturer’s instructions.

Mapping the transcriptional start sites of sub-genomic RNAs

Total RNA extracted from cambial scrapings of canes from GLRaV-1-infected

Chardonnay and Pinot Noir grapevines was used as a template to synthesize the first-strand cDNA using gene-specific primers (Table 3). The cDNA was ‘dC’-tailed as described above for determining the 5’end of the genome. The dC-tailed cDNA was then used in PCR using gene- specific nested primers and adapter primers (AAP and AUAP) supplied with the 5’RACE kit.

44 Conditions for PCR assays were the same as described above for 5’RACE. PCR products were cloned in pCR®2.1-TOPO® vector (ThermoFischer Scientific, Grand Island, New York).

Fifteen independent clones per amplicon were sequenced in both orientations at Retrogen Inc.

(San Diego, California). The sequences were aligned to the genome sequence of GLRaV-1 isolates WA-CH and WA-PN to identify transcriptional start sites (TSS) and determine the size of leader sequence for candidate sgRNAs. The dA-tailing method described above was used to resolve ambiguity of the 5’ terminal nucleotide for CPd1, CPd2 and p21 sgRNAs (Karasev et al.,

1995).

Sequence and secondary structure analysis

Nucleotide sequences generated in this study were edited, contigs assembled and sequence identities determined using Vector NTI Advance 11 sequence analysis software

(ThermoFischer Scientific, Grand Island, New York). Nt and amino acid (aa) sequences were aligned using CLUSTAL W multiple sequence alignment (Thompson et al., 1994). Phylogenetic relationships were inferred by employing the Neighbor-Joining (NJ) algorithm with 1,000 bootstrap replications using the Molecular Evolutionary Genetic Analysis software (MEGA) version 5 (Tamura et al., 2011). Secondary structure analysis of GLRaV-1 sequences was carried out using MFOLD software (Zuker et al., 1992).

45 Co-ordinates based on GLRaV- Primer 1 Amplicon IDa Sequence (5’ to 3’) WA-CH (nt) size (Kb) VA-F GTATTGTCCTAGTAGGTA 1 to 18 VA-R GTGGTTCGCACGCGAGTCAGGGTT 5449 to 5472 ~ 5.4 VB-F ATATGCGGAAGGAGAAGTGCAGG 5014 to 5036 VB-R TCGACCACCCGTAATTGGGGAGAA 9050 to 9073 ~ 4.0 VC-F GTGACGTTTTGTTGCAGC 7420 to 7437 VC-R GAGAAAGTTGCAATGCAGCACGGA 13908 to 13931 ~ 6.5 VD-F TAGATTGTTCTCAGCGACAGCCGT 13850 to 13873 VD-R GGGTGAGATATATAAATAAACGGAAACG 18704 to 18731 ~ 4.8

Table 1. List of primers used to amplify overlapping genome segments of GLRaV-1. Primer IDs,

primer sequence in 5’ to 3’ orientation and size of the PCR amplicons are shown. Co-ordinates

are based on the genome sequence of GLRaV-1 WA-CH isolate. aF and R represent forward and reverse primers, respectively.

Co-ordinates based Primer on GLRaV-1 WA- ID Sequence (5’ to 3’) CH (nt) 5'-OPa CAAGAAGACTTGATTTCCAT 882 to 901 5'-IP1a GCGTGCTTCACTTTATAGTATCCCTCCTC 649 to 677 5'-IP2a TCGTTTCCTTTCACTCAGGGACGA 372 to 395 3'-OPa CGCTGTTGGCC/TGAAGACAGTC 18142 to 18162 3'-IP1a GTAGCCCGAAAGATG 18166 to 18180 3'-IP2a ACGCTAATTACCGGGGAGT 18613 to 18631 AAPb GGCCACGCGTCGACTAGTACGGGIIGGGIIGGGIIG - AUAPb GGCCACGCGTCGACTAGTAC - mAAPc GGCCACGCGTCGACTAGTACTTTTTTTTTTTTTTTT -

46 Table 2. List of primers used to determine the 5’- and 3’- terminal sequences of GLRaV-1

genome using 5’RACE kit (ThermoFischer Scientific, Grand Island, New York). Primer co-

ordinates are based on GLRaV-1 WA-CH isolate. aOP is gene-specific outer primer, IP1 and IP2 are gene-specific nested internal primers. bAAP

and AUAP are abridged anchor primer and abridged universal anchor primer, respectively,

supplied with the 5’RACE kit. cmAAP is the modified AAP primer with oligo dT.

Primer Co-ordinates based on GLRaV-1 ID Sequence (5’ to 3’) WA-CH (nt) CP-OPa AATCTTTACCAACCCCGAGATGA 13484 to 13506 CP-IP1a TGGCTAGACGTGAAATGGGCGATA 13432 to 13455 CP-IP2a AAGACACAGTACTCGAAGCCAGCA 13275 to 13298 P21-OPa AACCAGGTGTAAAGAAGCTCACCG 17304 to 17327 P21-IP1a ATGCATACAGGGAAGGCGTCGATA 17258 to 17281 P21-IP2a GACTTTAACAGCGACGA 17120 to 17136 P24-OPa CGAAGCGAACGTCCTGAAAGACAT 17915 to 17938 P24-IP1a AGATGCACGGAATGTACTTACGCG 17893 to 17916 P24-IP2a TGAATTCTCGGTGGTT 17752 to 17769

Table 3. List of primers used to map the 5’ terminus of the sgRNAs specific to CP, CPd1, CPd2,

p21 and p24 from WA-CH and WA-PN isolates. aOP is gene-specific outer primer, IP1 and IP2 are gene-specific nested internal primers.

47 RESULTS

Sequence-dependent approach for sequencing of GLRaV-1 genome

The genome of GLRaV-1 isolates WA-CH and WA-PN was amplified in four overlapping cDNA fragments using sequence-specific primers (Fig. 4B). The derived cDNA clones specific to each fragment were sequenced in both orientations by ‘primer walking’ using progressive sequence-specific primers. Sequences generated from cDNA clones and 5’- and 3’- terminal sequences were assembled to build the full-length genome sequence. The genome of

GLRaV-1 thus determined was 18,731 nt in length for the WA-CH isolate and 18,946 nt in length for the WA-PN isolate. GenBank accession numbers for WA-CH and WA-PN are

KU674796 and KU674797, respectively. The genome of these two isolates is larger than the

18,659 nt reported for the Canadian isolate (NCBI accession no. NC_016509). However, it should be noted that nucleotide sequence of the Canadian isolate was obtained using Illumina

HiSeq and assembled by CLC genomic workbench v. 4.8 and the 5’- and 3’-terminal sequences were not authenticated.

The genome of WA-CH and WA-PN isolates encode nine predicted ORFs and the overall genome organization is similar to that of GLRaV-1 previously reported from Australia and

Canada. Individual ORFs encoded by these isolates are similar in size with the number of nt and aa in each ORF, sequentially from the 5’end of the genome, as follows: ORF1a (6612/2204),

ORF1b (1581/527), p7 (177/59), HSP70h (1629/543), p55 (1446/482), CP (966/322), CPd1

(1500/500), CPd2 (1320/440), p21 (567/189) and p24 (630/210). The nt and aa sequence identity of individual ORFs encoded by WA-CH and WA-PN isolates were compared among themselves and with corresponding sequences of GLRaV-1 from Australia and Canada (Table 4). The results indicated more than 95% identity at nt and aa sequence level between WA-CH and WA-PN

48 isolates compared with respective sequences of GLRaV-1 from Canada and Australia. A notable difference was observed in length of the intergenic region (IGR) between ORF 1b and ORF 2 among the four isolates. The IGR of WA-CH and WA-PN isolates are 770 nt and 965 nt in length compared to the corresponding sequence of 754 nt and 796 nt in Canadian and Australian isolates. The large size IGR in WA-PN isolate is due to the presence of a 183 nt sequence duplication separated by two nucleotides between the repeats. The two sequence repeats have an identity of 94%.

The ORF1a and ORF1b of WA-CH and WA-PN isolates shared greater than 95% nt and aa sequence identity between them and with corresponding ORF1a and ORF1b sequences of the

Canadian isolate (Table 4). No such comparisons were made with the Australian isolate due to the availability of partial sequence of ORF1a. However, the ORF1a sequence of the Australian isolate showed about 82 percent identity at the nt level and about 90 percent identity at the aa level with corresponding ORF1b sequence in WA-CH, WA-PN and Canadian isolates. All other

ORFs encoded by WA-CH and WA-PN isolates showed identities between 75 and 90 percent with their homologs in both Canadian and Australian isolates, except the CP and CPd1. The CP in the case of Canadian isolate and CP and CPd1 in the case of Australian isolate showed above

90% identity at aa level with WA-CH and WA-PN isolates. These results suggest non-uniform sequence identity across the genome of GLRaV-1 isolates currently sequenced. Overall, ORF- by-ORF comparisons indicate that GLRaV-1 isolates from Washington are more closely related to each other than either of them to the virus isolates reported from Canada and Australia.

The ORF1a, with an estimated molecular mass of 244 kDa, contains one papain-like leader protease (L-Pro) domain between aa 297-371. The conserved catalytic Cysteine (Cys) and

Histidine (His) residues, characteristic of papain-like proteases in positive-stranded RNA viruses

49 (Gorbalenya et al., 1991; Peng et al., 2001), are present in the L-Pro domain (Fig. 5). The MET domain was putatively mapped to aa 451-741, the HEL domain to aa 1899-2164 and the AlkB domain to aa 1466-1651. Like in GLRaV-1 and 4, the AlkB domain is located between MET and

HEL domains. The MET and HEL domains contain the DxxR and GKT motifs that are hallmark of positive strand RNA viruses (Gorbalenya et al., 1990; Rozanov et al., 1992; Koonin et al.,

1993). A comparison of the homologs of AlkB, a member of the 2OG-Fe(II) oxygenase superfamily (Aravind and Koonin, 2001; van den Born et al., 2008), among the grapevine- infecting ampeloviruses showed the presence of a conserved ‘AlkB core domain’ with high aa sequence identity (45%) to that of GLRaV-3 (Fig. 6). A signature ‘HAD’ motif is present in both

GLRaV-1 and -3 compared to ‘HKD’ motif in the AlkB domain of GLRaV-4 and its strains -5, -

6, -9, -Pr and -Car. The ORF1b has a 53-nt overlap with ORF1a in WA-CH and WA-PN isolates similar to the Canadian and Australian isolates. ORF1b is likely produced by +1 ribosomal frameshift as predicted for other closteroviruses (Agranovsky et al., 1994; Ling et al., 2004). An alignment of the overlapping sequence between ORF1a/1b overlap indicated a conserved hexanucleotide ‘UUUCGA’ around +1 ribosomal frameshift region in GLRaV-3 and GLRaV-1 isolates (Fazeli and Rezaian, 2000). In both viruses, the frameshift mechanism involves a ribosomal slippage at +1 P-site of tRNAPhe resulting in translation of amino acid Phenyl alanine in two frames, UUU and UUC (Fig. 7).

50

* * GLRaV-2_L1 486 --VIYPDGRCYLAHMRYLCAFYCRPFRESDYALGMWPTVARLR-ACVEKNFGV--EACGIALRGYYTSRNVYHCDYDSAYVKYFRNLSGRIGG-- 573 GLRaV-2_L2 760 --IRYPNGFCYLAHCRYACAFLLRGFDPKRFDIGAFPTAAKLR-NRMVSELGE--RSLGLNLYGAYTSRGVFHCDYDAKFIKDLRLMSAVIAG-- 847 GLRaV-1WA-CH 297 ELYYGDKYWCWLQ------LAVLNGNNLLAGSFESCISVRKLKRMLRFN-----VKLEKTEEAN---IFH--VGNKPTVSLAEVDDRC---- 368 GLRaV-1WA-PN 297 ELYYGDKYWCWLQ------LAVLNGNNLLAGSFESCISVRKLKRMLRFN-----VKLEKTEEAN---IFH--VGNKPTVSLAEVDDRC---- 368 GLRaV-1CAN 297 ELFYGDKYWCWLQ------LAVLNGNNLLAGSFESCISVRKLKRMLRFN-----VKLEKTDEAN---IFH--VGNKPTVSLAEVDDRC---- 368 GLRaV-6 435 -NFRGDRGFCWLP------LYTSSDIPVSQYPTGGLVRLFALYDKFG----PVPIVKSGKY-----YH--YDPKGRRHTKFPNVWVGAE- 506 GLRaV-3 302 AFVCTTKGWCWFN------NERLRGEIYRRRCFSSSFSIG-FLMHLGFR----SLKVIRFAGTN---ILH-----MPSLNEERTFGWKGGD- 371 GLRaV-4 302 -NWRGDRGFCWLP------LYLNSELPLTQYPTGGLVRLFTLYDKFG----PVPIVKSGKY-----YH--YDVKGKKHIKFPNVWVGAQ- 371 GLRaV-5 331 -NFRGDRGFCWLP------LYTSSDIPISQYPTGGLVRLFALYDKFG----PVPIVKSGKY-----YH--YDPKGRKHIKFPNVWVGAE- 401 GLRaV-7 497 KFSSVKIGYCWLD------AFNLSKKAIPDWLVPLPFIPLYLIFKCGVPKTILRMIRRTGNG---VCHFDRRYAHAHSIAKMTDLLGA-- 575 GLRaV-9 337 -NWRGDKGFCWLP------LYTASDIPISQYPTGGLVRLFTLYDKFG----PVPIVKSGKY-----YH--YDVKGKKHIKFPNVWVGAAP 408

Figure 5. Comparison of L-protease domain among GLRaVs. Multiple sequence alignment of L-protease domain from different

GLRaVs showing conserved nucleotides (shaded regions), and catalytic cysteine (C) and histidine (H) residues (shown by asterisk) of

L-protease family. L1 and L2 are tandem leader protease domains. Numbers at the ends of alignment indicate the position of amino 51

acid residues in the respective protein sequence. Viruses listed here and corresponding accession numbers are: Grapevine leafroll-

associated virus 1 (GLRaV-1) isolates WA-CH (KU674796), WA-PN (KU674797) and Canada (CAN; NC_016509), Grapevine

leafroll-associated virus 2 (GLRaV-2; JX559644), Grapevine leafroll-associated virus 3 (GLRaV-3; NC_004667), Grapevine leafroll

associated virus 4 (GLRaV-4; FJ467503), Grapevine leafroll-associated virus 5 (GLRaV-5; NC_016081), Grapevine leafroll-

associated virus 6 (GLRaV-6; FJ467504), Grapevine leafroll-associated virus 7 (GLRaV-7; NC_016436) and Grapevine leafroll-

associated virus 9 (GLRaV-9; AY297819).

GLRaV-1WA-CH 1466 ---KSSAIIKDAIPVKVEVSATRDAQTAKEREKFKGARKLVCSDIFV------DKLRGREVAFFSKCSR 1525 GLRaV-1WA-PN 1466 ---KSSAIIKEAIPVKVEESATRDAQTATEPERFKGARKLVCSDIFV------DKLRGREVAFFSKCSR 1525 GLRaV-1CAN 1466 ---KSSAIIEDVTPVKVEVSAPRNVQTAMEREKFKGARKLVSSDIFV------DKLRGRDVAFYSKYSR 1525 GLRaV-3 1492 KAIAHMVEKKQVQAEPPKQRNLTIDENKANKQLCMFRKCSCGVQLDVYNEATIATRFSNVFTFVDNLKGRSAVFFSKLGE 1571 GLRaV-4 1612 ---HETVFVEEPTCETDTTSDSSEQPIEEVSCEESHLTCSCGIDINVK-PFTVPAPLP--LIGGDKLNGREAWFYSRKGD 1685 GLRaV-5 1612 ---HETVFVEEPTCETDTTSDSSEQPIEEVSCEESHLTCSCGIDINVK-PFTVPAPLP--LIGGDKLNGREAWFYSRKGD 1685 GLRaV-6 1609 ---HETVFVEEPMCETDTTSDSSEKSIEEVSCEESHLTCSCGIDISVR-PFTVPAPLP--LIGGDKLNGREAWFCSRNGD 1682 GLRaV-9 1312 ---CEPTVEADNVSVADTVSDSSEQSMEELMPEIKHLKCDCGVDINVR-HFTVPGSLS--LVNGDRLNGREAWFYSRNGD 1385 GLRaV-Pr 1561 ---PSGVITVEKTAAPEADPNAVTMVEHMESDSFSPLKCACGIEIPVD-RFVSPGPLP--LLRGDSMNNREAWFYSRGGE 1634 GLRaV-Car 1562 ---SREIEVSEPKVTTQTTRPSTP------PRVCRLSCQCGVIIPVR-PFKAPGDLP--LVSGDVMRNRSAWFYSRGGE 1628

* * * * * ** GLRaV-1WA-CH 1526 RYVYNGGSHASQGWNKALDELREELKL-DESYDHCLIQKYRKGATIGFHADDEKCYTSGVSVVTVNLNGQARFRVRSNKT 1604 GLRaV-1WA-PN 1526 GYVYNGGSHASQGWNKALDELREELKL-DESYDHCLIQKYRKGATIGFHADDEKCYTPGVSVVTVNLNGQARFRVRSNKT 1604 GLRaV-1CAN 1526 RYVYNGGSHASQGWNKALDELREELKL-DESYDHCLIQKYRKGATIGFHADDEKCYTSGVSVVTVNLNGQARFRVRSNKT 1604 GLRaV-3 1572 GYTYNGGSHVSSGWPRVLEDILTAIKY-PSVFDHCLVQKYKMGGGVPFHADDEECYPSDNPILTVNLVGKANFSIKCRKG 1650 GLRaV-4 1686 GYSYVGGSHVSRGWLNILNRYISNTGLNPNLFDHCLIQKYECGAGIPYHKDNEPVYPKNNPILTIHVSGEGMFSIRCNNG 1765 GLRaV-5 1686 GYSYVGGSHVSRGWLNILNRYISNTGLNPNLFDHCLIQKYECGAGIPYHKDNEPVYPKNNPILTIHVSGEGMFSIRCNNG 1765 GLRaV-6 1683 PYSYVGGSHNSRGWPNVLNKYIVNTGLNPATFNHCLIQRYVAGAGIPYHKDNEAVYPKNNPILTIHVSGEGMFSIRCYDG 1762 GLRaV-9 1386 SYSYVGGSHESRGWPNILNRFVAATGLNPAMFNHCLVQRYKAGAGIPYHKDNEPVYPKNNPILTIHVSGEGMFSVRCLNS 1465 52 GLRaV-Pr 1635 GYSYTGYSHKSRGWLSILDRFVSATGLKSSMFDHCLIQKYNRGAGIPFHKDNEPVYPIGNPILTIHLSGEGMFSIKCGTG 1714

GLRaV-Car 1629 SYSYTGGSHKSRGWLDILDAYISACGLEPELFDHCLIQKYQPDSGLNFHKDDEPVYPRMNPVLKIHASGTGVFSVCCNEG 1708

* * * * * GLRaV-1WA-CH 1605 GEVVEHLLGDGDVFVMSPGMQRDFKHSVESLDEGRVSITLRNATVDY 1651 GLRaV-1WA-PN 1605 GEVVEHLLGDGDVFVMSPGMQRDFKHSVESLDEGRVSITLRNATVDY 1651 GLRaV-1CAN 1605 GEIVEHLLGDGDVFVMSPGMQRDFKHSVESLDEGRVSITLRNATVDY 1651 GLRaV-3 1651 GKVMVMNVASGDYFLMPCGFQRTHLHSVSSIDEGRISLTFRATRRVF 1697 GLRaV-4 1766 SGGVLLKPPS--WFLMPFGFQITHQHSVTCA-TVRVSMTFRSTEVIT 1809 GLRaV-5 1766 SGGVLLKPPS--WFLMPFGFQITHQHSVTCA-TVRVSMTFRSTEVIT 1809 GLRaV-6 1763 SGKLPMKEPC--WFMMPFGFQVSHQHSVACA-TVRVSMTFRSTEVIK 1806 GLRaV-9 1466 SGEIMLKEPC--WFLMPFGFQVSHQHSVTCA-TVRVSMTFRSTEVIK 1509 GLRaV-Pr 1715 CGELLMTKPC--WFLMPCGFQKTHLHSVTCS-SERVSLTFRATQQLK 1758 GLRaV-Car 1709 SGQLDMTDPC--YFCMPNGFQISHYHAVRCT-TERVSLTFRSTKFVE 1752

Figure 6. Comparison of AlkB domain among GLRaVs. The solid line shows an AlkB core domain (adapted from van den Born et al.,

2008), which has high sequence homology within viral AlkB proteins. Dark shaded regions in the alignment represent identical amino

acids and the light shaded regions indicate similar amino acids. Asterisks mark the conserved amino acids in AlkB family (Aravind

and Koonin 2001). Numbers at the ends of alignment indicate the position of amino acid residues in the respective protein sequence.

Viruses listed here and corresponding accession numbers are: Grapevine leafroll-associated virus 1 (GLRaV-1) isolates WA-CH

(KU674796), WA-PN (KU674797) and Canada (CAN; NC_016509), Grapevine leafroll-associated virus 2 (GLRaV-2; JX559644),

Grapevine leafroll-associated virus 3 (GLRaV-3; NC_004667), Grapevine leafroll associated virus 4 (GLRaV-4; FJ467503),

Grapevine leafroll-associated virus 5 (GLRaV-5; NC_016081), Grapevine leafroll-associated virus 6 (GLRaV-6; FJ467504),

Grapevine leafroll-associated virus 7 (GLRaV-7; NC_016436), Grapevine leafroll-associated virus 9 (GLRaV-9; AY297819),

Grapevine leafroll-associated virus Pr (GLRaV-Pr; NC_011702), Grapevine leafroll-associated virus Car (GRLaV-Car; FJ907331).

53

GCTGGAAAAGTGAGTGACGTTTTGTTGCAGCAATTGCAGCGGAACGATCGCTTTCGAGTCGATTGA S D V L L Q Q L Q R N D R F F E S I E GTGACGTTTTGTTGCAGCAATTGCAGCGGAACGATCGCTTTCGAGTCGATTGAGTGTTTCTCGGCT

Figure 7. Nucleotide sequence showing the ORF1a/1b overlap and +1 ribosomal frameshift region in GLRaV-1. ORF1a sequence is

shown at the top and ORF 1b sequence is shown at the bottom. One letter translation products are shown in between the nucleotide

sequences. Underlined sequence shows the 53 nt overlap between ORF1a-1b. +1 ribosomal frame shift region (as reported by Fazeli

and Rezaian, 2000) is shown in bold. Stop codon on ORF 1a is represented in bold, italicized letters.

3' 5' NTR ORF 1a ORF 1b p7 HSP70h p55 CP CPd1 CPd2 p21 p24 NTR WA-CH vs 97.02 98.3 97.22 97.92 96.55 97.32 97.67 95.99 97.72 96.84 WA-PN 93.7 (97.19) (98.86) (94.92) (98.34) (95.23) (98.14) (98.4) (94.09) (98.41) (95.24) 93.92 WA-CH vs 94.48 96.01 83.33 83.88 81.78 86.58 84.83 78.61 79.3 81.2 CAN 87.84* (95.19) (96.91) (88.14) (88.95) (84.02) (95.34) (88.4) (74.55) (79.89) (84.29) 71.55* WA-CH vs 79* 82.13 91.67 90.99 88.61 91.23 90.62 86.24 88.95 88.78 AUS (91.71*) (90.89) (89.83) (93.37) (88.38) (92.86) (91.8) (84.09) (86.77) (86.67) 74.72* WA-PN vs 94.68 96.2 83.89 83.7 82.19 86.58 84.63 78.84 79.12 80.73 CAN 86.43* (95.09) (97.49) (89.83) (88.21) (82.99) (95.03) (88.2) (74.32) (80.42) (81.9) 79.32* WA-PN vs 79* 82.01 91.11 90.44 87.92 91.23 90.55 86.17 89.3 88.31 54 AUS (91.96*) (91.27) (88.14) (92.63) (87.34) (93.17) (91.8) (84.09) (88.36) (86.19) 73.33* CAN vs 79* 82 84 85 82 86 84 80 79 79 AUS (92*) (90) (86) (89) (83) (93) (87) (75) (76) (80) 68

Table 4. Comparison of nucleotide and amino acid sequence identity of ORFs between GLRaV-1 isolates. Amino acid sequence

identity levels are shown in parenthesis. *Comparison of percent nucleotide/amino acid sequence identity with partial sequence of

Australian or Canadian isolate available in GenBank.

Sequence variation in the 5’- and 3’-NTRs

The 5’NTR of WA-CH and WA-PN isolates is 857 nt and 922 nt in length, respectively.

The first nt at the 5’terminus, determined by 5’RACE, is guanine (G), similar to that in BYV,

LIYV, GLRaV-7, and -4 and its strain -6 (Agranovsky et al., 1994; Klaassen et al., 1995;

Ghanem-Sabanadzovic et al., 2012; Jelkmann et al., 2012). Like the 5’NTR of other closteroviruses, the 5’NTR sequences of both isolates are rich in A + U content (63%). A pairwise comparison of the 5’NTR sequences of WA-CH and WA-PN isolates with the Canadian isolate indicate that the latter is 11 nt shorter, which could be due to the lack of verification of the terminus by 5’RACE. Nevertheless, a comparison among the three isolates revealed some commonalities as well as unique differences in their 5’NTR sequences. In WA-CH, WA-PN and

Canadian isolates, four 65 nt repeats were identified with sequence identities ranging from 53% to 92% between the repeats. These repeats are present between positions 181 nt to 440 nt in WA-

CH, 181 nt to 440 nt in WA-PN and 105 nt to 371 nt in Canadian isolate. In addition, an extra repeat of 65 nt sequence was identified in the 5’NTR of WA-PN isolate between 441 nt and 494 nt with 50% to 78% sequence identity to the four repeats in WA-PN (Fig. 8).

In order to verify if the observed repeats in the 5’NTR are not due to errors during RT-

PCR, cloning or sequencing, additional experiments were conducted using samples from ornamental grapevine cultivars (V. vinifera, cv. Purpurea, V. californicum, cv. Roger’s Red) tested positive for GLRaV-1 (Alabi et al., 2011). Extracts from petioles were used in RT-PCR to amplify approximately 900 nt fragment covering the complete 5’NTR using primers VA-F

(5’GTATTGTCCTAGTAGGTA 3’, corresponding to nt 1 to 18 in WA-CH or WA-PN) and

5’OP (5’ CAAGAAGACTTGATTTCCAT 3’, complementary to nt 882 nt to 901 of WA-CH isolate, Tables 1 and 2). Analysis of sequences from 10 independent clones confirmed the

55

presence of the four 65 nt repeats in WA-CH and five 65 nt repeats in WA-PN.

A comparison of the secondary structures predicted by the MFOLD program showed that the 5’NTRs of both isolates can be folded into a complex secondary structure consisting of a long Stem-Loop (SL) structure with several sub-structural hairpin loops of variable lengths (Fig.

9).

The 3’NTR of WA-CH and WA-PN isolates was determined to be 456 nt and 410 nt, respectively. The difference in the length was due to an extra 44 nt from positions 18,548 nt to

18,591 nt in the genome of WA-CH isolate (Fig. 10). In order to confirm this difference, virus sequences were amplified from two ornamental grape samples listed above with primers 3’NTR-

F (5’ GTAGCCCGAAAGATG 3’, corresponding to positions 18165 nt – 18180 nt in WA-CH) and VD-R (5’ GGGTGAGATATATAAATAAACGGAAACG 3’, complimentary to positions

18704 – 18731 nt). Analysis of sequences from 10 independent clones confirmed presence of the extra sequence in WA-CH. The 3’NTR of GLRaV-1 isolates from Canada and Australia was reported to be 468 nt and 360 nt, respectively. A 23 nt sequence reported at the 3’ terminus of the

Canadian isolate between 18,637 nt and 18,659 nt was not observed in the 3’NTR sequences of

WA-CH and WA-PN isolates. An attempt to amplify the 3’ terminal sequence of GLRaV-1 isolates using primer sequence 5’ GTAGCCCGAAAGATG 3’, corresponding to positions 18165 nt – 18180 nt in WA-CH isolate, and the 23 nt sequence, complementary to 18,637 nt and 18,659 nt of the Canadian isolate, failed to yield the approximately 490 nt amplicon expected from virus-infected samples (data not shown). This anomaly suggests that the 3’terminal 23 nt sequence reported in the Canadian isolate may be an artifact and do not constitute an integral part of the 3’-terminus of GLRaV-1. The 3’ NTR of Australian isolate is shorter in length compared to the 3’NTR sequences from Canadian and Washington isolates. The difference might be due to

56

mispairing of oligo dT primer to the 3’-poly(A)-tailed dsRNA, resulting in shorter 3’NTR in case of the Australian isolate. Similar to the 5’NTR, the predicted secondary structure of 3’NTRs of

WA-CH and WA-PN isolates showed complex structures consisting of several SL structures

(Fig. 11).

A comparison of the size of 5’ and 3’NTRs of GLRaV-1 isolates WA-CH and WA-PN with the size of corresponding 5’ and 3’ terminal sequences of members of the family

Closteroviridae is shown in Table 5. The data indicate that GLRaV-1 isolates WA-CH and WA-

PN have unusually long non-translated sequences at their 5’ and 3’ terminus and this could be a characteristic of GLRaV-1 isolates.

57

Genus Virus 5'NTR 3'NTR GenBank accession number Ampelovirus GLRaV-1 857 - 922 410 - 456 KU674796, KU674797 JQ423939, KM058745, GU983863, JQ796828, JX559645, 672 - 802 259 - 450 GLRaV-3 KJ174518, GQ352631, GQ352633 GLRaV-4 216 - 218 127 - 129 FJ467503, KJ810572, NC_016416 GLRaV-4-like 213 - 218 127 - 130 NC_011702, NC_016417, FJ467504, KJ810572, viruses PMWaV-1 353 129-131 NC_010178, AF414119, KJ872494 PBNSPaV 301-302 231-247 KC590347, KC590345, KC590344, KJ792852, NC_009992 BVBaV 781 335 NC_022072 Closterovirus GLRaV-2 105 - 106 206 - 215 NC_007448, KF220376 BYV 107 166 – 182 AF056575, NC_001598, X73476, AF190581 CTV 106 – 109 267 - 362 GQ454870, EU857538 CYLV 324 227 NC_013007

58 SCFaV 227 215 NC_008366 Crinivirus LIYV (RNA 1) 97 218 NC_003617 LIYV (RNA 2) 692 191 NC_003618 BYDV (RNA 1) 83 249 NC_010560 BYDV (RNA 2) 511 465 NC_010561 LCV (RNA 1) 72 226 NC_012909 LCV (RNA 2) 268 98 NC_012910 ToCV (RNA 1) 93 225 NC_013258 ToCV (RNA 2) 584 179 NC_013259 SPaV (RNA 1) 264 197 NC_005895 SPaV (RNA 2) 240 186 NC_005896 Velarivirus GLRaV-7 47 192 NC_016436, HE588185 LChV-1 76 207 NC_001836 CoV-1 67 269 HM588723

Table 5. Comparison of the length of 5’ and 3’ NTRs among viruses in family Closteroviridae. Viruses listed in the table are as

follows: Genus Ampelovirus: Grapevine leafroll-associated virus 1 (GLRaV-1), Grapevine leafroll-associated virus 3 (GLRaV-3),

Grapevine leafroll associated virus 4 (GLRaV-4) and GLRaV-4 like viruses: Grapevine leafroll-associated virus 5 (GLRaV-5),

Grapevine leafroll-associated virus 6 (GLRaV-6), Grapevine leafroll-associated virus 9 (GLRaV-9), Grapevine leafroll-associated

virus Pr (GLRaV-Pr), Grapevine leafroll-associated virus Car (GRLaV-Car), Pineapple mealybug wilt-associated virus 1 (PMWaV-

1), Plum bark necrosis stem pitting-associated virus (PBNSPaV), Blackberry vein banding associated virus (BVBaV). Genus

Closterovirus: Grapevine leafroll-associated virus 2 (GLRaV-2), Beet yellows virus (BYV), Citrus tristeza virus (CTV), Carrot

yellow leaf virus (CYLV), Strawberry chlorotic fleck-associated virus (SCFaV). Genus Crinivirus: Lettuce infectious yellows virus

59 (LIYV), Bean yellow disorder virus (BYDV), Lettuce chlorosis virus (LCV), Tomato chlorosis virus (ToCV); Strawberry pallidosis-

associated virus (SPaV); genus Velarivirus: Grapevine leafroll-associated virus 7 (GLRaV-7), Little cherry virus 1 (LChV-1),

Cordyline virus 1 (CoV-1). Abbreviations of the viruses and corresponding accession numbers are listed in the table.

CAN 1 ------GTAGGTATCGAACCAGACAATCCCAATCCTACCATTGCTTTCCATTTCAGATCTGAAATCCACACTTCTTCTCTTCTCTCCGCTTCCGAGCATTCTTAC------99 WA-CH 1 GTATTGTCCTAGTAGGTATCGAACCAGACAATTCCAATCCTACCATTGCTTTCCATTCCAGATCTGAAATCCACACTTCTTCTCTTCTCTCTGCTTCCGAGCATTCTTCCTCACTTTCCT 120 WA-PN 1 GTATTGTCCTAGTAGGTATCGAACCAGACAATCCCAATCCTACCATTGCTTTCCATTCCAGAACTGAAATCCACACTTCTTCTCTTCTCTCTGCTTCCGAGCATTCTTTCTCACTTTCCT 120 ********************* ************************ **** **************************** **************** *

CAN 100 ------TCTCATTCCTTTTTCTCTCGAGTGCTAAAACCACTCCCTTCTCACTACCCTTTTACCTCTCCCTT 164 WA-CH 121 GTTCAGACACGGCGTGCTGTATAACGGTTCGTTTGTCTTTTCTCGTCTCTCACTTTCTTATTCCTTTTTCTTTCGAGTGCTAATACCACTCCCTTCTCACCACCCTCTTACATCTGCCTT 240 WA-PN 121 GTTCAGACACGGCGTGCTGTATAACGGTTCGTTTGTCTTTTCTCGTCTCTCACTTTCTTATTCCTTTTTCTCTCGAGTGCTAATACCACTCCCTTCTCACCACTCTCTTACATCTCCCTT 240 *** ************ *********** **************** ** ** **** *** ****

CAN 165 TTCGTTTCCTTTCTCTCAGGGACGAAGTATATACTTCACCTCTCTACTGTTCTACTTCTTCTCCTTTTCTTTCCTTTTTCTCTTGAGTGCTGATATCACTCTATTCTCACTACTTTTTCA 284 WA-CH 241 TTCGTTTCCTTTCTTTCAGGGACGAAGCACATACTTCATCTCTTTATTCTTCTACTTCTTCTCCTCTTCTTTCCTTTTTCTCTTGAGTGCTAATATCACTCCCTTCTTACCACTTTTTCA 360 WA-PN 241 TTCGTTTCCTTTCTTTCAGGGACGAAGCACACACTTCATCTCTCTACTCTCTTACTTCTTCTTCTTTTCTTTCCTTTTTCTCTCGAGTGCTAATACCACTCCCTTCTCACCACTCTCTTA 360 ************** ************ * * ****** **** ** * * ********** ** ***************** ******* *** ***** **** ** *** * * *

CAN 285 CTTCTCCCTTTTTGTTTCCTTTCATTCAGGGACG-AAGCACGTACTTCTTCTCTTTACTCTTCTCCCTTTTCTCCTTATCT------364 WA-CH 361 CTTCTCCCTTTTCGTTTCCTTTCACTCAGGGACG-AAGCACTAGCTTCTTCTCTTTACTCTTCTACCTCTTCTCCTTTTCT------440 WA-PN 361 CATCTCTTCTTTTCTTTCCTTTTTCTCTTGAGTGCAAATATCACTCCCTTCTTACCACTTTT-TCATTTCTCCCTTTTCGTTTCCTTTCACTCAGGGACTGAGCACCTGCTTCTTCTCTTT 480 * **** *** ******** ** * * ** * ***** *** ** * * ** * ** *

CAN 365 ------TTCCTTTCTTTTTTATTTTTTATTTAACTCTATTTATCCGCCTTCTTTCTTTTCGTATCGTTCATTTCCTTCTCTTTTCTTTTTCTTTTTATCGT 459 WA-CH 441 ------TTCCTTTCTTTTTTATTTTCTATTATATTTTATTTATCCGCCTTCTCTCTTTTCGTACCGATCATTTCCTTTTCTTTTCTTTCTCCTTTTAAATT 535 WA-PN 481 ACTTTTCTACCTCTTCTCCTTTTCTTTCCTTTCTTTTTCATTTTCTACTTAATTTTATTTATCCGCCTTCTTTCTTTTCGTACCAATCATTTCCTTCTCTTTTCTTTTTCTTTTTAACTT 600 ************* ***** ** * * * **************** ********** * ********** ********** ** ***** *

CAN 460 TACTTTTTCGCCTTCTTTCTTATTGTATCCATCGTTTCCTTTTCTTTTCTCCTTAGTTCGTCACTCTAACGTTTTCCCGAACCACTCCATTTCTTTTCATTTCCTTTCATTTCGCGTGCT 579 WA-CH 536 TACTTTTTCACCTTCTTTCTTTTTGTATCTATCGTTTCCTTTTCTTTTCTTCATAGTTCGTCACTCATACGTTTTCTCGTTTTCCTCCTTTTCTTAAGTTTTCCTTTCATTTCGCGTGCT 655 WA-PN 601 TACCTATTCACCCCTTTTCTTTTTGTATCCATCGTTTCCTTTTCTTTTCTTCATAGTTCGTCACTCATACGTTTTCCCGTATTTCTCCTTTTCTTAATATTTCCTTTTATTTCGCGTGTT 720 *** * *** ** ****** ******* ******************** * ************* ******** ** **** ****** ******** ********** *

CAN 580 TCACTTTATTGTTTCCCCTTTCTTTTACTGTCTAGGTTCTAATTTTTCCTTTTCCTCCTTAGTATTTTGCAGTATCGGTTTCATTTCTAACTTTTTTATTCGTCTTGTACTTCCTTTTAT 699 WA-CH 656 TCACTTTATAGTATCCCTCCTCTTTTACCGTCTAGGTTTTACCTTTTCCTTTTCCTCTTTAGCATTTTGTTGTTTCGGTTTCATTTTTTACATATTTTTCCGTCTTGCGTTTCCTTTTAT 775 WA-PN 721 TCACTTTATAGTATCCCTCCTCTTTTACTGTCTAGGTTTTATCTTTTCCTTTTTTCTTTTAGCATTTTGCTGTTTCGGTTTCGGTTTTTACGTATTTTTTCGTCTAGCGTTTCCTTTTAT 840 60 ********* ** **** ******** ********* ** ********** **** ****** ** ******** ** * ** * *** * ***** * **********

CAN 700 CTTCGCTTTTCTCGTTTTTGTCGTTCCCCATTGTTCTCTTTTCTTATTTTTATATTTCCTTTCTTCTCGTTTATTTAAAATC 781 WA-CH 776 CTTCGTTTTTCTTGTTTTTGTCGGTCCCTTTTGTTCTCTTTTCTTTTCTCTATTTTTCCTTTCTTCTCGCTTATTTAAAATC 857 WA-PN 841 CTTCGTTTTTCTTGTCTTTGTCGGTCCCTTTTGTTCTTTTTTCTTTTTGCTATTTTTCCTTTCTCCTAGCTTATTTAAAATC 922 ***** ****** ** ******* **** ******* ******* * *** ********** ** * ************

Figure 8. Multiple sequence alignment of 5’NTRs from GLRaV-1 isolates WA-CH, WA-PN and Canada. The 11 nt sequence

identified by 5’RACE in WA-CH and WA-PN at the 5’-terminus is underlined. Bold and italicized nucleotides represent the 65 nt

sequence present in WA-CH and WA-PN. Pink, blue, green and orange colored text represents the four 65 nt repeats present in

GLRaV-1 isolates WA-CH, WA-PN and Canada. An extra 65 nt sequence present in WA-PN is shown in purple colored text.

A B

Figure 9. MFOLD predicted secondary structures from the 5’NTR sequence of GLRaV-1 isolates WA-CH (A) and WA-PN (B).

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Figure 10. Multiple sequence alignment of 3’NTR sequence from GLRaV-1 isolates WA-CH,

WA-PN, Australia and Canada. Distinct differences in the 3’NTR sequence are marked in different font styles. Numbers at the ends of alignment indicate the position of nucleotide residue in the full length sequence of respective GLRaV-1 isolates.

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A B

Figure 11. MFOLD predicted secondary structures from the 3’NTR sequence of GLRaV-1 isolates WA-CH (A) and WA-PN (B).

Discrimination of GLRaV-1 isolates based on sequence variability in the 5’NTR

The 5’NTR sequences of WA-CH and WA-PN were used to examine the potential of molecular-based typing of GLRaV-1 variants in a vineyard. For this purpose, random samples were collected from Pinot noir vines showing GLD symptoms in a commercial vineyard and tested by RT-PCR using diagnostic primers described earlier (Alabi et al., 2011). Samples from vines tested positive for GLRaV-1 were subsequently used in RT-PCR to amplify the 5’NTR with primers VA-F (5’ GTATTGTCCTAGTAGGTA 3’) and 5’-OP (5’

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CAAGAAGACTTGATTTCCAT 3’; Tables 1 and 2). The resulting amplicons from nine independent vines were cloned and the derived sequences compared with 5’NTR sequences of

WA-CH and WA-PN isolates. A phylogenetic analysis of these sequences using the Neighbor- joining algorithm showed three clades, with six sequences clustering with WA-CH to form group

1, another two sequences branching separately to form group 2 and one sequence clustering with

WA-PN to form group 3 (Fig. 12A). Multiple sequence alignment showed 93 percent identity between group 1 and 2 sequences, whereas sequences in group 3 showed 82 percent identity with group 1 and 2. Following an in silico analysis of 5’NTR sequences, XhoI restriction enzyme was found to be a reliable marker for developing a molecular typing method, based on RT-PCR-

RFLP profiles of 5’NTR sequences, to quickly discriminate GLRaV-1 isolates in vineyards. The

DNA fragments amplified in RT-PCR using primers VA-F and 5’-OP were digested individually with XhoI and resolved by agarose gel electrophoresis. The results showed unique restriction patterns distinguishing all sequences into three groups (Fig. 12B; Table 6). Sequences in group 2 gave two bands of approximately 650 bp and 200 bp sizes due to the presence of one restriction site for XhoI. Sequences in group 3 gave three bands of approximately 550 bp, 250 bp and 130 bp size due to the presence of two restriction enzyme sites. Conversely, the amplicons from group 1 isolates remained as a single band due to the absence of the restriction site for XhoI.

These results indicate that the phylogenetic tree based on the 5’NTR sequences is in congruence with RFLP analysis. Based on these results, it can be concluded that RT-PCR-RFLP profiling of

5’NTR sequences is a relatively quick assay for detecting variants of the virus in vineyards.

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A.

B.

Figure 12. RT-PCR based RFLP method for detecting GLRaV-1 variants in Washington vineyards. (A) Neighbor-joining phylogenetic tree showing clustering of GLRaV-1 variants into three groups. Boot strap values are indicated at the branch nodes. The bar shown below the tree indicates 0.005 substitutions per site. (B) Agarose gel electrophoresis of PCR products after restriction digestion with XhoI. M: 1kb plus DNA ladder (ThermoFischer Scientific, Grand

Island, New York); lanes 1, 2 and 3 shows RFLP digestion pattern for GLRaV-1 variant groups

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1, 2 and 3, respectively, corresponding to the three groups shown in the tree. Arrows indicate the size of DNA molecular marker on the left and individual fragments released upon restriction digestion on the right. Specific DNA bands released after restriction digestion are shown by asterisks.

Number of fragments after restriction GLRaV-1 variant Sample ID digestion group WA-PN 3 3 PN17-3 3 3 PN18-3 2 2 PN60-2 2 2 WA-CH 1 1 PN17-2 1 1 PN17-6 1 1 PN16-3 1 1 PN19-1 1 1 PN8-1 1 1 PN8-2 1 1

Table 6. Restriction fragment length polymorphism (RFLP) analysis of the 5’NTR sequence from various GLRaV-1 isolates in Washington. List of the samples, fragments generated upon restriction digestion and corresponding variant groups are listed.

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Expression of 3’-coterminal sgRNAs and identification of transcription start sites

By analogy with other closteroviruses, ORFs downstream of the RGB would be expected to be expressed via 3’-coterminal sgRNAs. Using riboprobe specific to the 3’-end of GLRaV-1 genome, Fazeli and Rezaian (2000) previously observed three major 3’-coterminal sgRNA species in Northern blot analysis of dsRNA isolated from grapevines infected with GLRaV-1.

These sgRNAs were putatively identified as specific to CP, p21 and p24. To extend these studies, total RNA was isolated from cambial scrapings of canes collected from virus-positive grapevines and analyzed by Northern blot hybridization with riboprobes corresponding p24, the

3’ most ORF in GLRaV-1 genome. As shown in Fig. 13, three sgRNAs were detected at higher levels and were putatively designated as sgRNAs specific to CP, p21 and p24, respectively.

Among them, the sgRNA corresponding to p24 gene accumulated at the highest level, followed by sgRNAs for CP and p21, respectively. Two barely visible bands between CP and p21 were tentatively identified as sgRNAs corresponding to CPd1 and CPd2. The other poorly visible bands above the CP sgRNA band were tentatively designated as sgRNAs corresponding to p55,

HSP70h/p7 and genomic RNA. Overall, these results support previous observations (Fazeli and

Ali Rezaian, 2000) that 3’-coterminal sgRNAs accumulate at variable amounts indicating that

GLRaV-1-encoded sgRNAs are expressed at different levels and/or exhibit variable turnover rates in infected grapevine tissues.

Additional experiments were conducted to determine the 5′-terminal sequence of sgRNAs and examine their leader sequences. Using total RNA isolated from virus-infected grapevine samples, the 5’ ends of CP, CPd1, CPd2, p21 and p24 sgRNAs were amplified by the 5’RACE system using a combination of gene-specific and an abridged anchor primer and the amplicons were cloned and nucleotide sequence determined. The derived sequences were aligned with the

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genome of GLRaV-1 to map the exact nucleotide located at the 5’-end of individual sgRNAs. By this approach, the location of transcription start site (TSS) for CP, CPd1, CPd2, p21 and p24 was mapped, respectively, to 13,064, 14,096, 15,613, 16,981 and 17,625 nt on the genome of

GLRaV-1 isolate WA-CH. The 5’-end sequences for sgRNAs p7, HSP70h and p55 could not be identified, despite several attempts. This could be, in part, due to low abundance of these sgRNAs in virus-infected grapevine samples used in this study (Jarugula et al., 2010). The 5′ terminal nt of these five sgRNAs were found to be purines and conserved in the genome of WA-

CH, WA-PN, Australian and Canadian isolates. Using the location of TSS with respect to the genome sequence of WA-CH isolate, the size of sgRNAs for CPd1, CPd2, p21 and p24 were estimated as 4,699 nt, 2,226 nt, 1,744 nt and 1,234 nt, respectively. The size of CP sgRNA for

WA-CH and WA-PN isolates was 5,668 nt and 5,623 nt, respectively, due to difference in the

IGR length between p55 and CP among WA-CH (95 nt IGR) and WA-PN (96 nt IGR). Using the location of TSS and the putative start codon of each ORF, the length of leader sequence for CP,

CPd1, CPd2, p21 and p24 ORFs was determined to be 67/68, 27, 15, 49 and 18nt, respectively

(Table 7). The 67/68, 49 and 18 nt leader sequences corresponding to CP, p21 and p24 sgRNAs are located entirely in the intergenic region (IGR) upstream of their respective initiation codons.

In contrast, the leader sequences of CPd1 and CPd2 were mapped within CP and CPd1 ORFs, respectively. This is due to 22 nt and 3 nt IGR for CPd1 and CPd2, respectively, compared to 95 nt, 79 nt and 43 nt IGR corresponding to CP, p21 and p24. Beside these differences in TSS of the

CP sgRNA, the leader sequence of sgRNAs for CPd1, CPd2, p21 and p24 are identical between the WA-CH and WA-PN.

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Size of Transcriptional start site of the subgenomic Size of leader Subgenomic subgenomic RNAs on the RNA (WA-CH in sequence WA-CH RNA genome of WA-CH (nt) nt) and WA-PN (nt) CP 13,064 5,668 67 and 68* CPd1 14,096 4,636 26 CPd2 15,613 3,119 15 p21 16,981 1,751 49 p24 17,625 1,107 18

Table 7. Characteristics of the sub-genomic RNAs specific to five ORFs encoded by the genome of WA-CH and WA-PN isolates. * Size of CP leader sequence on the genome of WA-PN isolate

Figure 13. Northern blot hybridization of GLRaV-1 RNA using riboprobe specific to p24. Arrow head indicates putative sgRNA bands identified in the blot. Asterisks mark the tentative location of GLRaV-1 sgRNAs.

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DISCUSSION

In this study, the genome sequence of GLRaV-1 was determined using tiled, overlapping amplicons spanning the entire genome. In addition, the rapid amplification of cDNA ends

(RACE) method was used for confirmation of the 3' and 5' NTR terminal sequences. By these approaches, the complete genome of two isolates of GLRaV-1 was determined to be 18,731 nt

(WA-CH) and 18,946 nt (WA-PN). The sequence data indicated that GLRaV-1 has the second largest positive strand ssRNA genome among the monopartite members of the family

Closteroviridae, after CTV that has ~19.3 kb genome (Karasev et al., 1995). Relative to the genome size of 18.5–18.7 kb in GLRaV-3 isolates (Maree et al., 2013) and 18.6 kb in Blackberry vein banding-associated virus (BVBaV, Thekke-Veetil et al., 2013), the 18.7 to 18.9 kb size of

GLRaV-1 makes it the largest genome among members of the genus Ampelovirus. It is interesting to note that these viruses with unusually large genome size have been documented only in perennial crops, although the significance of this observation needs further research.

Nevertheless, the complete genome sequence of GLRaV-1 generated in this study, together with the complete genome sequences determined previously for GLRaV-2, -3, -4 and its strains, and

GLRaV-7 (Zhu et al., 1998; Ling et al., 2004; Maliogka et al., 2009; Ghanem-Sabanadzovic et al., 2010, 2012; Jelkmann et al., 2012; Thompson et al., 2012; Velasco et al., 2015), should provide opportunities for elucidating the comparative molecular biology of grapevine-infecting members of the family Closteroviridae.

The overall genome organization of GLRaV-1 isolates WA-CH and WA-PN with hallmark gene array consisting of nine ORFs corresponds to that of GLRaV-1 sequences reported from Australia and Canada (Fig. 4). Like other monopartite members in the genus

Closterovirus and Ampelovirus, GLRaV-1 isolates WA-CH and WA-PN encodes two

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overlapping ORFs (ORF1a/b) that makeup the replication-associated gene module, with ORF1a containing putative domains for a papain-like proteinase, methyltransferase, AlkB, and helicase and ORF1b having a RdRp domain. The six conserved motifs in the helicases (Gorbalenya and

Koonin, 1993) and eight conserved sequence motifs in the polymerases (Koonin et al., 1993) of positive-stranded RNA viruses of superfamily I are observed in the helicase and polymerase domain of GLRaV-1 isolates WA-CH, WA-PN and Australia isolates (Fazeli and Rezaian,

2000). Comparative genome sequence analysis indicated that GLRaV-1 uses +1 ribosomal frameshift mechanism to express ORF1b (Agranovsky et al., 1994). Isolates of GLRaV-1 has a

53 nt overlap between ORF1a and ORF1b. In this context, a 58-113 nt overlap was also reported between ORF1a and ORF1b of GLRaV-3 (NC_004667, Ling et al., 1998; EU344893, Engel et al., 2008; EU259806, Maree et al., 2008; GQ352633, GQ352632 and GQ352631, Jooste et al.,

2010; GU983863, Jarugula et al., 2010; JQ655296 and JQ655295, Bester et al., 2012; JQ796828,

Seah et al., 2012; JQ423939, Fei et al., 2013; KM058745, Bester et al., 2014). In contrast to

GLRaV-1 and GLRaV-3, the putative start codon for ORF1b is located -5 to -3 relative to stop codon of the ORF1a in GLRaV-4 and its strains (Maliogka et al., 2009; Ghanem-Sabanadzovic et al., 2010, 2012; Thomson et al., 2012; Velasco et al., 2015). A comparison of +1 frameshifting among closterovirids shows that the slippery sequence involved in translational frameshifting in

GLRaV-1 (UUUC) is identical to GLRaV-3 and BVBaV, and distinct from other closteroviruses such as BYV, GLRaV-4, GLRaV-6 (GUUU; Agranovsky et al., 1994; Ghanem- Sabanadzovic et al., 2012), CTV (GUUCGG; Karasev et al., 1995), GLRaV-2 and GLRaV-7 (UAG; Zhu et al.,

1998; Jelkmann et al., 2012) and LIYV (AAAG; Klaassen et al., 1995). Outside the family

Closteroviridae, the +1 ribosomal frameshift in GLRaV-1 is similar to the frameshift region

(UCC UUU CGU) in chronic bee paralysis virus that infects insects, fijiviruses and

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amalgamaviruses that infect plants and influenza A virus that infect vertebrates (Firth et al.,

2012; Firth, 2014).

The first five ORFs of the remaining eight ORFs, downstream of the RGB, comprise the quintuple gene block and the remaining three ORFs towards the 3’end of the genome seems to be unique to GLRaV-1. In analogy with their homologs in other members of the family

Closteroviridae (Agranovsky et al., 1995; Tian et al., 1999; Peremyslov et al., 1999, 2004;

Satyanarayana et al., 2000, 2004; Alzhanova et al., 2001, 2007; Reed et al., 2003; Lu et al.,

2004), many of these proteins are likely to be multifunctional involved in various aspects of the virus life cycle, including cell-to-cell and systemic movement and suppressors of RNA silencing to counter host antiviral responses. Although the occurrence of a diverged duplicate of the CP, designated as CPm, is a common feature with many closteroviruses, the presence of two diverged copies of the CPm, CPd1 and CPd2, only in the genome of GLRaV-1 is an additional unique feature of the virus in the family Closteroviridae. It has been speculated that such gene duplication is a consequence of ancient duplication events occurred during the evolution and diversification of closteroviruses (Dolja et al., 2006). In analogy with BYV (Peremyslov et al.,

2004) and CTV (Satyanarayana et al., 2004), GLRaV-1 is likely to have a bipolar architecture, with CP encapsidating most of the filamentous particles and CPm forming the main component of the virion ‘head’. However, it remains to be examined whether one or both of CPd1 and CPd2 is a component of the segmented ‘head’ of GLRaV-1 virions.

The most striking feature of GLRaV-1 is the large size 5’ NTR, varying between 857 nt and 922 nt, compared to other members of the family Closteroviridae (Table 5). It should be noted that incomplete data on the 5’ and 3’ terminal sequences of the Canadian and Australian isolates precluded comparative analysis of the noncoding region sequences among the four

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GLRaV-1 isolates. Irrespective, the presence of an unusually long 5’NTR appears to be a characteristic feature of ampeloviruses (Table 5). To our knowledge, no other plant virus appears to have such a long and variable noncoding region at the 5’terminus of the genome. A comparison with members of the genus Ampelovirus shows that the 5’NTR of GLRaV-1 isolates sequenced in this study is longer than those reported for other viruses, such as BVBaV and

GLRaV-3 (Table 5). Within the family Closteroviridae, members of the genus Ampelovirus have a long 5’NTR relative to other monopartite members in the genera Closterovirus and Velarivirus.

Among the bipartite members of the genus Crinivirus, RNA 2 has a long 5’NTR compared to

RNA 1, except SPaV that has more or less similar size 5’NTR sequence in both RNA segments.

These results further supports the grouping of GLRaV-1 with BVBaV and GLRaV-3 as a sister clade, designated subgroup II, within the genus Ampelovirus (Maliogka et al., 2008). Outside the family Closteroviridae, the wheat-infecting Triticum mosaic virus (TriMV) in the family

Potyviridae, was reported to have a 739 nt 5’NTR (Tatineni et al., 2009a). Outside plant viruses, long 5’NTRs between 650 nt and 1300 nt has been reported in animal/human-infecting viruses in the family Picornaviridae (reviewed in Jackson and Kaminski, 1995; Belsham and Sonenberg,

1996). Unlike these viruses, the 5’NTR sequence of GLRaV-1 isolates contain no AUG codons preceding the translation initiation codon of the ORF1a/b. From a practical point of view, the

RFLP analysis of the 5’NTR (Fig. 12) can offer finer discrimination of GLRaV-1 variants circulating in a vineyard. Together with genetic diversity studies reported earlier (Kominek et al.,

2005; Alabi et al., 2011, Fan et al., 2015), knowledge of variability in the 5’NTR could enhance the power of discriminating GLRaV-1 isolates in epidemiological studies and to infer evolutionary relationships among genetically divergent variants of the virus.

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Our analysis detected sequence duplications in the 5’NTR, similar to previous reports with the 5’NTR of GLRaV-3 (Jarugula et al., 2010). The most striking feature noted for GLRaV-

1 is the presence of four repeats of 65 nt sequence arranged end-to-end in WA-CH, WA-PN and

Canada isolates. In WA-PN isolate, an extra 65 nt sequence is present downstream of the four end-to-end repeats. Although these repeats in both GLRaV-1 and -3 appear to be of similar size, their sequences are highly dissimilar between GLRaV-1 and -3. To the best of our knowledge, the 65 nt repeats in the 5’-nontranslated regions have not been reported in other plant viruses.

The significance of ~65 nt sequence repeats in GLRaV-1 and -3 and the lack of conservation between these sequences remains obscure. It is possible that these repeats in 5’NTRs of both viruses could serve to provide a backup function, in terms of playing a regulatory role in translation and replication. Nevertheless, the exact mechanism of the sequence repeats in the 5’ noncoding regions of GLRaV-1 and -3 and their role in gene regulation and expression as well as biological fitness advantage beneficial to these viruses need further investigations.

In contrast, long sequence duplications were not observed in the noncoding regions at the

3’-terminus of GLRaV-1 and -3. Outside the family Closteroviridae, a unique direct repeat sequence of 32 nt and 76 nt was reported in the 3’NTR of PVY-N (Robaglia et al., 1989) and an isolate of Pepper mottle virus (Warren and Murphy, 2003).

Similar to GLRaV-3 (Jarugula et al., 2010), both 5’ and 3’NTRs of GLRaV-1 have the potential to form complex secondary structures. However, very little is known about the relationship of primary sequences and predicted secondary structures of the NTRs of these viruses. An approximately 100 to 120 nucleotide residues at the 5’terminus is highly conserved between variants of each virus, implying that these conserved sequences harbor important requirements for virus replication. Preliminary reports with GLRaV-3 have indeed suggested that

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sequences at the beginning of the 5’NTR contains critical element(s) required for virus replication (Jarugula et al., 2012). In the case of CTV, the 5’NTR forms two stable SL structures, despite having high sequence diversity (López et al., 1998) and these conserved secondary structures were shown to be essential for replication and assembly (Gowda et al., 2003). The highly conserved 3’NTR of CTV isolates could be folded into a complex secondary structure composed of 10 SLs and this complex structure was shown to harbor critical elements for efficient replication of the virus (Satyanarayana et al., 2002). It can be hypothesized that sequences in the 5’ and 3’ NTRs of GLRaV-1 could be harboring functional elements and their functions could be mediated by a combination of linear sequence and structural elements. The development of an amenable reverse genetic system for GLRaV-1 would help to examine the important and diverse roles of 5’ and 3’ NTRs in the virus life cycle.

Characterization of the sgRNA profiles by Northern blot hybridization revealed that three of the eight 3' co-terminal sgRNAs were present at higher levels in GLRaV-1-infected grapevine

(Fig. 5). An analogous set of sgRNAs corresponding to CP, p21 and p24 have also been reported by Fazeli and Ali Rezaian (2000). These results suggest that synthesis of 3’-coterminal sgRNAs occurs by complex gene expression strategy that is characteristic of the members of the family

Closteroviridae. Among the three sgRNAs, the sgRNA specific to the p24 showed higher accumulation followed by the sgRNAs specific to CP and p21, suggesting that genes located nearer to the 3’end of the genome are usually expressed at higher levels as reported with other closteroviruses (Navas-Castillo et al., 1997; Hagiwara et al., 1999). The difference in relative amounts of the three different sgRNAs and the inability to detect sgRNAs corresponding to other sgRNAs under our experimental conditions could be due to their differential expression in infected leaves. Such asymmetric accumulation of the individual sgRNAs could be due to

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temporal and quantitative regulation of their expression in virus-infected plants, as reported with other closteroviruses such as CTV (Navas-Castillo et al., 1997), BYV (Hagiwara et al., 1999),

Lettuce infectious yellows virus (LIYV) (Yeh et al., 2000), Sweet potato chlorotic stunt virus

(SPCSV) (Kreuze et al., 2002) and GLRaV-3 (Jarugula et al., 2010).

To further study the transcriptional strategy of GLRaV-1 sgRNAs, the 5’-termini of the sgRNAs specific to CP, CPd1, CPd2, p21 and p24 ORFs were mapped relative to the genomic

RNA. Repeated attempts were failed to identify the 5’-termini of the sgRNAs specific to p7,

HSP70h and p55 and this could likely be due to low level of expression of these sgRNAs in samples collected for analysis. The 5’ terminus of the sgRNAs specific to CP and p24 is adenylate, and CPd1, CPd2 and p21 is guanylate, similar to the 5’ terminus of the genomic RNA.

Variability in the 5’terminal nucleotide of the sgRNAs was also observed with SPCSV in the genus Crinivirus (Kreuze et al., 2002) and BYV in the genus Closterovirus (Agranovsky et al.,

1994; Peremyslov and Dolja, 2002; Vitushkina et al., 2007). In contrast, all sgRNAs in CTV

(Ayllón et al., 2003) and sgRNAs of GLRaV-3 characterized so far (Jarugula et al., 2010) have adenylate at the 5’terminus. The AU-rich leader sequences of sgRNAs do not have any conserved sequence motifs.

In summary, this study provided the complete genomic sequence of GLRaV-1 and revealed a number of new features characteristic of the virus. Based on the sequence data, it appears that the genome of GLRaV-1 is the second largest, after CTV, among members of the family Closteroviridae. Another striking novel feature of the virus is the long 5’ and 3’ NTRs and the presence of 65 nt sequence repeats in the 5’NTRs compared to other closterviruses.

Together with the information generated on 3’-coterminal sgRNAs, this study provided a

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foundation for the development of an amenable reverse genetic system in the future to conduct a more detailed analysis of the virus gene functions.

ACKNOWLEDGEMENTS

This research was supported, in part, by the WSU Agricultural Research Center, the Wine

Advisory Committee, the Washington Wine Commission, Washington State Grape & Wine

Research Program, Northwest Center for Small Fruits Research, and Altria - Chateau Ste.

Michelle Wine Estates. Thanks to Dr. Sridhar Jarugula for contributing to Northern blotting experiments and Dr. Siddarame Gowda for critical review.

AFFLIATION OF CO-AUTHORS

Dr. Sridhar Jarugula

Ohio Agriculture Research and Development Center, Ohio State University, Wooster, Ohio,

United States of America.

Dr. Rayapati A. Naidu

Department of Plant Pathology, Washington State University, Irrigated Agriculture Research and

Extension center, Prosser, Washington, United States of America.

AUTHORS' CONTRIBUTIONS

Conceived and designed the experiments: BP, SJ and RAN. Performed the experiments:

BP and SJ. Analyzed the data: BP and RAN. Contributed reagents/materials/analysis tools:

RAN. Wrote the paper: BP and RAN.

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REFERENCES

Agranovsky, A. A., Koonin, E. V., Boyko, V. P., Maiss, E., Frötschl, R., Lunina, N. A., and Atabekov, J. G. 1994. Beet yellows closterovirus: complete genome structure and identification of a leader papain-like thiol protease. Virology. 198:311–324. doi:10.1006/viro.1994.1034.

Agranovsky, A. A., Lesemann, D. E., Maiss, E., Hull, R., and Atabekov, J. G. 1995. “Rattlesnake” structure of a filamentous plant RNA virus built of two capsid proteins. Proc. Natl. Acad. Sci. U.S.A. 92:2470–2473. doi:10.1073/pnas.92.7.2470.

Alabi, O. J., Al Rwahnih, M., Karthikeyan, G., Poojari, S., Fuchs, M., Rowhani, A., and Naidu, R. A. 2011. Grapevine leafroll-associated virus 1 occurs as genetically diverse populations. Phytopathol. 101:1446–56. doi:10.1094/PHYTO-04-11-0114.

Alabi, O. J., Gutha, L. R., Casassa, L. F., Harbertson, J., Mirales, M., Beaver, C. W., Davenport, J., and Naidu, R. A. 2012. Impacts of grapevine leafroll disease on own- rooted wine grape cultivar in cool climate conditions. 63rd American Society for Enology and Viticulture National Conference. Portland, OR, U.S.A pp. 69-70.

Almeida, R. P. P., Daane, K. M., Bell, V. A., Blaisdell, G. K., Cooper, M. L., Herrbach, E., and Pieterson, G. 2013. Ecology and management of grapevine leafroll disease. Front. Microbiol. 4:94. doi:10.3389/fmicb.2013.00094.

Alkowni, R., Zhang, Y. P., Rowhani, A., Uyemoto, J. K., and Minafra, A. 2011. Biological, molecular, and serological studies of a novel strain of grapevine leafroll-associated virus 2. Virus Genes 43:102-110. doi:10.1007/s11262-011-0607-7.

Al Rwahnih, M., Dolja, V. V., Daubert, S., Koonin, E. V., and Rowhani, A. 2012. Genomic and biological analysis of grapevine leafroll-associated virus 7 reveals a possible new genus within the family Closteroviridae. Virus Res. 163:302-309. doi:10.1016/j.virusres.2011.10.018.

Alzhanova, D. V., Napuli, A. J., Creamer, R., and Dolja, V. V. 2001. Cell-to-cell movement and assembly of a plant closterovirus: roles for the capsid proteins and HSP70 homolog. EMBO J. 20:6997–7007. doi:10.1093/emboj/20.24.6997.

Alzhanova, D. V., Prokhnevsky, A. I., Peremyslov, V. V., and Dolja, V. V. 2007. Virion tails of Beet yellows virus: coordinated assembly by three structural proteins.Virology 359:220–26. doi:10.1016/j.virol.2006.09.007.

Aravind, L., and Koonin, E. V. 2001. The DNA-repair protein AlkB, EGL-9, and leprecan define new families of 2-oxoglutarate- and iron-dependent dioxygenases. Genome Biol. 2:research0007.1-research0007.8

Ayllón, M. A., Gowda, S., Satyanarayana, T., Karasev, A. V., Adkins, S., Mawassi, M., Guerri, J., Moreno, P., and Dawson, W. O. 2003. Effects of modification of the transcription initiation site context on citrus tristeza virus subgenomic RNA synthesis. J. Virol. 77:9232–9243.

78

Bahder, B. W., Poojari, S., Alabi, O. J., Naidu, R. A., and Walsh, D. B. 2013. Pseudococcus maritimus (Hemiptera: Pseudococcidae) and Parthenolecanium corni (Hemiptera: Coccidae) are capable of transmitting grapevine leafroll-associated virus 3 between Vitis x labruscana and Vitis vinifera. Environ. Entomol. 42:1292–1298. doi:10.1603/EN13060.

Basso, M. F., Fajardo, T. V. M., Santos, H. P., Guerra, C. C., Ayub, R. A., and Nickel, O. 2010. Leaf physiology and enologic grape quality of virus-infected plants. Trop. Plant Pathol. 35:351– 59. doi:10.1590/S1982-56762010000600003.

Belsham, G. J., and Sonenberg, N. 1996. RNA–protein interactions in regulation of picornavirus RNA translation. Microbiol. Rev. 60:499-511.

Benson, G. 1999. Tandem repeats finder: A program to analyze DNA sequences. Nucleic Acids Res. 27:573–580. doi:10.1093/nar/27.2.573.

Bertazzon, N., Borgo, M., Vanin, S., and Angelini, E. 2010. Genetic variability and pathological properties of grapevine leafroll-associated virus 2 isolates. Eur. J. Plant Pathol. 127:185-197. doi:10.1007/s10658-010-9583-3.

Bester, R., Maree, H. J., and Burger, J. T. 2012. Complete nucleotide sequence of a new strain of grapevine leafroll-associated virus 3 in South Africa. Arch. Virol.157:1815-19. doi:10.1007/s00705-012-1333-8.

Bester, R., Pepler, P. T., Burger, J. T., and Maree, H. J. 2014. Relative quantitation goes viral: An RT-qPCR assay for a grapevine virus. J. Virol. Methods 210:67-75. doi:10.1016/j.jviromet.2014.09.022.

Dolja, V. V., Kreuze, J. F., and Valkonen, J. P. T. 2006. Comparative and functional genomics of closteroviruses. Virus Res. 117:38–51. doi:10.1016/j.virusres.2006.02.002.

Donda, B. P., and Naidu, R. A. 2015. Sequence analysis of grapevine leafroll-associated virus 1 from Washignton vineyards. Page 26 in: American Phytopathological Society (APS) annual meeting, Pasadena, California, USA.

Engel, E. A., Girardi, C., Escobar, P. F., Arredondo, V., Dominguez, C., Perez-Acle, T., and Valenzuela, P. D. 2008. Genome analysis and detection of a Chilean isolate of Grapevine leafroll associated virus-3. Virus Genes 37:110-118. doi: 10.1007/s11262-008-0241-1.

Fan, X., Hong, N., Dong, Y., Ma, Y., Zhang, Z. P., Ren, F., Hu, G., Zhou, J., and Wang, G. 2015. Genetic diversity and recombination analysis of grapevine leafroll-associated virus 1 from China. Arch. Virol. 160:1669-1678. doi:10.1007/s00705-015-2437-8.

Fazeli, C. F., and Rezaian, A. M. 2000. Nucleotide sequence and organization of ten open reading frames in the genome of grapevine leafroll-associated virus 1 and identification of three subgenomic RNAs. J. Gen. Virol. 81:605–615. doi:10.1099/0022-1317-81-3-605.

79

Fei, F., Lyu, M-D., Li, J., Fan, Z-F., and Cheng, Y-Q. 2013. Complete nucleotide sequence of a Chinese isolate of reveals a 5’UTR of 802 nucleotides. Virus genes 46:182-185. doi:10.1007/ s11262-012-0823-9.

Firth, A. E., Jagger, B. W., Wise, H. M., Nelson, C. C., Parsawar, K., Wills, N. M., Napthine, S., Taubenberger, J. K., Digard, P., and Atkins, J. F. 2012. Ribosomal frameshifting used in influenza A virus expression occurs within the sequence UCC UUU CGU and is in the +1 direction. Open Biol. 2:120109. doi:10.1098/rsob.120109.

Firth, A. E. 2014. Mapping overlapping functional elements embedded within the protein-coding regions of RNA viruses. Nuc. Acids Res. 42:12425-12439. doi:10.1093/nar/gku981.

Ghanem-Sabanadzovic, N. A., Sabanadzovic, S., Uyemoto, J. K., Golino, D., Rowhani, A. 2010. A putative new ampelovirus associated with grapevine leafroll disease. Arch. Virol. 155:1871– 76. doi:10.1007/s00705-010-0773-2.

Ghanem-Sabanadzovic, N. A., Sabanadzovic, S., Gugerli, P., and Rowhani, A. 2012. Genome organization, serology and phylogeny of grapevine leafroll-associated viruses 4 and 6: Taxonomic implications. Virus Res. 163:120–128. doi:10.1016/j.virusres.2011.09.001.

Golino, D. A., Wolpert, J., Sim, S. T., Benz, J., Anderson, M., and Rowhani, A. 2009a. Virus effects on vine growth and fruit components of three California ‘Heritage’ clones of Cabernet Sauvignon. Pages 243-244 in: Proc. XVI Int. Counc. Study Viruses Virus-Like Dis. Grapevine. Dijon, France.

Golino, D. A., Wolpert, J., Sim, S. T., Benz, J., Anderson, M., and Rowhani, A. 2009b. Virus effects on vine growth and fruit components of Cabernet Sauvignon on six rootstocks. Pages 245-246 in: Extended Abstr. 16th Meet. ICVG. Dijon, France.

Gorbalenya, A. E., Koonin, E. V., and Wolf, Y. I. 1990. A new superfamily of putative NTP- binding domains encoded by genomes of small DNA and RNA viruses. FEBS Lett. 262:145– 148. doi:10.1016/0014-5793(90)80175-I.

Gorbalenya, A. E., Koonin, E. V., and Lai, M. M.-C. 1991. Putative papain-related thiol proteases of positive-strand RNA viruses Identification of rubi- and aphthovirus proteases and delineation of a novel conserved domain associated with proteases of rubi-, α- and coronaviruses. FEBS Lett. 288:201–205. doi:10.1016/0014-5793(91)81034-6.

Gorbalenya, A. E. & Koonin, E. V. 1993. Helicases: amino acid sequence comparisons and structure-function relationship. Curr. Opin. Struc. Biol. 3:419-429. doi:10.1016/S0959- 440X(05)80116-2.

Gowda, S., Satyanarayana, T., Ayllón, M. A., Moreno, P., Flores, R., and Dawson, W. O. 2003. The conserved structures of the 5′ nontranslated region of citrus tristeza virus are involved in replication and virion assembly. Virology. 317:50–64. doi:10.1016/j.virol.2003.08.018.

80

Habili, N., Komínek, P., and Little, A. 2007. Grapevine leafroll-associated virus 1 as a common grapevine pathogen. Plant Viruses 1:63-68. doi:10.1094/ PHYTO-04-11-0114.

Hagiwara, Y., Peremyslov, V. V., and Dolja, V. V. 1999. Regulation of closterovirus gene expression examined by insertion of a self-processing reporter and by northern hybridization. J. Virol. 73:7988–7993.

Jackson, R. J., and Kaminski, A. 1995. Internal initiation of translation in eukaryotes: The picornavirus paradigm and beyond. RNA 1:985-1000.

Jarugula, S., Gowda, S., Dawson, W. O., and Naidu, R. A. 2010. 3’-coterminal subgenomic RNAs and putative cis-acting elements of grapevine leafroll-associated virus 3 reveals “unique” features of gene expression strategy in the genus Ampelovirus. Virol. J. 7:180. doi:10.1186/1743- 422X-7-180.

Jarugula, S., Gowda, S., and Naidu, R. A. 2012. Nucleotide sequence at the beginning of the 5’ non-translated region is critical for replication of grapevine leafroll-associated virus 3. Extended Abstract. 17th Meeting of ICVG, Davis, CA, USA. pp 54-56.

Jelkmann, W., Fechtner, B., and Agranovsky, A. A. 1997. Complete genome structure and phylogenetic analysis of little cherry virus, a mealybug-transmissible closterovirus. J. Gen. Virol. 78:2067-2071. doi:10.1099/0022-1317-78-8-2067.

Jelkmann, W., Mikona, C., Turturo, C., Navarro, B., Rott, M. E., Menzel, W., Saldarelli, P., Minafra, A., and Martelli, G. P. 2012. Molecular characterization and taxonomy of grapevine leafroll-associated virus 7. Arch. Virol. 157:359–362. doi:10.1007/s00705-011-1176-8.

Jooste, A. E. C., Maree, H. J., Bellstedt, D. U., Goszczynski, D. E., Pietersen, G., and Burger, J. T. 2010. Three genetic grapevine leafroll-associated virus 3 variants identified from South African vineyards show high variability in their 5′UTR. Arch. Virol. 155:1997–2006. doi:10.1007/s00705-010-0793-y.

Karasev, A. V., Boyko, V. P., Gowda, S., Nikolaeva, O. V., Hilf, M. E., Koonin, C. L., Cline, N. K., Gumpf, D. J., Lee, R. F., Garnsey, S. M., Lewandowski, D. J., and Dawson, W. O. 1995. Complete sequence of the Citrus tristeza virus RNA genome. Virology 208:511-520. doi:10.1006/viro.1995.1182.

Klaassen, V. A., Boeshore, M. L., Koonin, E. V, Tian, T., and Falk, B. W. 1995. Genome structure and phylogenetic analysis of lettuce infectious yellows virus, a whitefly-transmitted, bipartite closterovirus. Virology. 208:99–110. doi:10.1006/viro.1995.1133.

Komar, V., Vigne, E., Demangeat, G., Lemaire, O., and Fuchs, M. 2010. Comparative performance analysis of virus-infected Vitis vinifera cv. Savagnin rose grafted onto three rootstocks. Am. J. Enol. Vitic.61:68–73.

81

Komínek, P., Glasa, M., and Bryxiová, M. 2005. Analysis of the molecular variability of grapevine leafroll-associated virus 1 reveals the presence of two distinct virus groups and their mixed occurrence in grapevines. Virus Genes 31:247-255. doi: 10.1007/s11262-005-3236-1.

Koonin, E. V., Dolja, V. V., and Morris, T. J. 1993. Evolution and taxonomy of positive-strand RNA viruses: implications of comparative analysis of amino acid sequences. Crit Rev Biochem Mol Biol 28:375– 430. doi:10.3109/10409239309078440.

Kreuze, J. F., Savenkov, E. I., and Valkonen, J. P. T. 2002. Complete Genome Sequence and Analyses of the Subgenomic RNAs of Sweet Potato Chlorotic Stunt Virus Reveal Several New Features for the Genus Crinivirus. J. Gen. Virol. 76:9260-9270. doi:10.1128/JVI.76.18.9260– 9270.2002

Langley, K. E., Vilareja, M. R., Fowler, A. V., Zamenhof, P. J., and Zabin, I. 1975. Molecular basis of beta-galactosidase alpha-complementation. Proc. Natl. Acad. Sci. USA. 72:1254-1257.

Lee, J., and Martin, R. R. 2009. Influence of grapevine leafroll associated viruses (GLRaV-2 and -3) on the fruit composition of Oregon Vitis vinifera L. cv. Pinot noir: phenolics. Food Chem.112:889–96. doi:10.1016/j.foodchem.2008.06.065.

Lee, J., Keller, K. E., Rennaker, C., and Martin, R. R. 2009. Influence of grapevine leafroll associated viruses (GLRaV-2 and -3) on the fruit composition of Oregon Vitis vinifera L. cv. Pinot noir: free amino acids, sugars, and organic acids. Food Chem.117:99–105. doi:10.1016/j.foodchem.2008.06.065.

Ling, K. S., Zhu, H. Y., Drong, R. F., Slightom, J. L., McFerson, J. R., and Gonsalves, D. 1998. Nucleotide sequence of the 3'-terminal two-thirds of the grapevine leafroll-associated virus-3 genome reveals a typical monopartite closterovirus. J. Gen. Virol. 79:1299-1307.

Ling, K. S., Zhu, H. Y., and Gonsalves, D. 2004. Complete nucleotide sequence and genome organization of grapevine leafroll-associated virus 3, type member of the genus Ampelovirus. J. Gen. Virol. 85:2099–2102. doi: 10.1099/vir.0.80007-0.

Little, A., Fazeli, C. F., and Rezaian, M. A. 2001. Hypervariable genes in Grapevine leafroll- associated virus 1. Virus Res. 80:109-116.

Little, A. 2004. Complete sequence, improved detection and functional analysis of grapevine leafroll-assicaited virus 1(GLRaV-1). PhD Thesis, The University of Adelaide, Australia, 101 pp.

López, C., Ayllón, M. A., Navas-Castillo, J., Guerri, J., Moreno, P., and Flores, R. 1998. Molecular variability of the 5′ and 3′ terminal regions of Citrus tristeza virus RNA. Phytopathol. 88:685-691. doi:10.1094/PHYTO.1998.88.7.685.

Lu, R., Folimonov, A., Shintaku, M., Li, W.-X., Falk, B. W., Dawson, W. O., and Ding, S.-W. 2004. Three distinct suppressors of RNA silencing encoded by a 20-kb viral RNA genome. Proc.

82

Natl. Acad. Sci. U.S.A. 101:15742–15747. doi:10.1073/pnas.0404940101.

Maliogka, V. I., Dovas, C. I., and Katis, N. I. 2008. Evolutionary relationships of virus species belonging to a distinct lineage within the Ampelovirus genus. Virus Res 135:125–135. doi:10.1016/j.virusres.2008.02.015.

Maliogka, V. I., Dovas, C. I., Lotos, L., Efthimiou, K., and Katis, N. I. 2009. Complete genome analysis and immunodetection of a member of a novel virus species belonging to the genus Ampelovirus. Arch. Virol. 154:209–218. doi:10.1007/s00705-008-0290-8.

Maree, H. J., Freeborough, M-J., Burger, J. T. 2008. Complete nucleotide sequence of a South African isolate of grapevine leafroll-associated virus 3 reveals a 5’UTR of 737 nucleotides. Arch. Virol. 153:755-657. doi:10.1007/s00705-008-0040-y.

Maree, H. J., Almeida, R. P. P., Bester, R., Chooi, K. M., Cohen, D., Dolja, V. V., Fuchs, M. F., Golino, D. A., Jooste, A. E. C., Martelli, G. P., Naidu, R. A., Rowhani, A., Saldarelli, P., and Burger, J. T. 2013. Grapevine leafroll-associated virus 3. Front. Microbiol. 82:1-21. doi:10.3389/fmicb.2013.00082.

Martelli, G. P., Agranovsky, A. A., Bar-Joseph, M., Boscia, D., Candresse, T., Coutts, R. H. A., Dolja, V. V., Falk, B. W., Gonsalves, D., Jelkmann, W., Karasev, A. V., Minafra, A., Namba, S., Vetten, H. J., Wisler, G. C., and Yoshikawa, N. 2002. ICTV study group on closteroviruses. The family Closteroviridae revised. Arch. Virol. 147:2039–44. doi:10.1007/s007050200048.

Martelli, G. P. 2012. Grape virology highlights: 2010-2012. Pages 13-31 in: Proc. 17th Congr. Int. Counc. Study Virus Virus-like Dis. Grapevine (ICVG), Davis, California, USA.

Martelli, G. P., Ghanem-sabanadzovic, N. A., Agranovsky, A. A., Rwahnih, M. Al, Dolja, V. V, and Dovas, C. I. 2012. Taxonomic revision of the family Closteroviridae with special reference to the grapevine leafroll-associated members of the genus Ampelovirus and the putative species unassigned to the family. J. Plant Pathol. 94:7–19. doi:10.4454/jpp.fa.2012.022.

Martelli, G. P. 2014. Directory of virus and virus-like diseases of the grapevine and their agents. J. Plant Pathol. 96(Suppl. 1):1–136. doi:10.4454/JPP.V96I1SUP.

Naidu, R. A., Rowhani, A., Fuchs, M., Golino, D., and Martelli, G. P. 2014. Grapevine leafroll: a complex viral disease affecting a high-value fruit crop. Plant Dis.98:1172–85. doi:10.1094/PDIS- 08-13-0880-FE.

Naidu, R. A., Maree, H. J., and Burger, J. T. 2015. Grapevine leafroll disease and associated viruses: A unique pathosystem. Annu. Rev. Phytopathol. 53:613-34. doi:10.1146/annurev-phyto- 102313-045946.

Navas-Castillo, J., Albiach-Martí, M. R., Gowda, S., Hilf, M. E., Garnsey, S. M., and Dawson, W. O. 1997. Kinetics of accumulation of citrus tristeza virus RNAs. Virology. 228:92–97. doi:10.1006/viro.1996.8369

83

Peng, C. W., Peremyslov, V. V, Mushegian, A. R., Dawson, W. O., and Dolja, V. V. 2001. Functional specialization and evolution of leader proteinases in the family Closteroviridae. J. Virol. 75:12153–12160. doi:10.1128/JVI.75.24.12153-12160.2001.

Peremyslov, V. V., Hagiwara, Y., and Dolja, V. V. 1999. HSP70 homolog functions in cell-to- cell movement of a plant virus. Proc. Natl. Acad. Sci. USA. 96:14771–76.

Peremyslov, V. V., and Dolja, V. V. 2002. Identification of the subgenomic mRNAs that encode 6-kDa movement protein and Hsp70 homolog of Beet yellows virus. Virology 295:299-306.

Peremyslov, V. V., Andreev, I. A., Prokhnevsky, A. I., Duncan, G. H., Taliansky, M. E., and Dolja, V. V. 2004. Complex molecular architecture of beet yellows virus particles. Proc. Natl. Acad. Sci. U. S. A. 101:5030–5035. doi:10.1073/pnas.0400303101.

Poojari, S., Alabi, O. J., and Naidu, R. A. 2013. Molecular characterization and impacts of a strain of grapevine leafroll-associated virus 2 causing asymptomatic infection in a wine grape cultivar. Virol. J. 10:324. doi:10.1186/1743-422X-10-324.

Rayapati, N. A., O’Neil, S., and Walsh, D. 2008. Grapevine Leafroll disease. WSU Extension Genetics 11:31-46.

Reed J.C., Kasschau, K.D., Prokhnevsky, A.I., Gopinath, K., Pogue, G.P., Carrington, J.C., and Dolja, V.V. 2003. Suppressor of RNA silencing encoded by Beet yellows virus. Virol. 306:203– 209. doi:10.1016/S0042-6822(02)00051-X.

Robaglia, C., Durand-Tardif, M., Tronchet, M., Boudazin, G., Astier-Manifacier, S., and Casse- Delbart, F. 1989. Nucleotide sequence of potato virus Y (N Strain) genomic RNA. J Gen Virol 70:935–947. doi:10.1099/0022-1317-70-4-935.

Rozanov, M. N., Koonin, E. V., and Gorbalenya, A. E. 1992. Conservation of the putative methyl transferase domain: a hallmark of the “Sindbis-like” supergroup of positive-strand RNA viruses. J. Gen. Virol. 73:2129–2134. doi:10.1099/0022-1317-73-8-2129.

Satyanarayana, T., Gowda, S., Mawassi, M., Albiach-Martí, M. R., Ayllón, M. A., and Robertson, C., Garnsey, S. M., and Dawson, W. O. 2000. Closterovirus encoded HSP70 homolog and p61 in addition to both coat proteins function in efficient virion assembly. Virology. 278:253–265. doi:10.1006/viro.2000.0638.

Satyanarayana, T., Gowda, S., Ayllón, M. A., Albiach-Martí, M. R., and Dawson, W. O. 2002. Mutational analysis of the replication signals in the 3’-nontranslated region of citrus tristeza virus. Virology. 300:140–152. doi:10.1006/viro.2002.1550.

Satyanarayana, T., Gowda, S., Ayllón, M. A., and Dawson, W. O. 2004. Closterovirus bipolar virion: evidence for initiation of assembly by minor coat protein and its restriction to the genomic RNA 5’ region. Proc. Natl. Acad. Sci. 101:799–804. doi:10.1073/pnas.0307747100.

84

Seah, Y. M., Sharma, A. M., Zhang, S., Almeida, R. P., and Duffy, S. 2012. A divergent variant of Grapevine leafroll-associated virus 3 is present in California. Virol. J. 9:235. doi:10.1186/1743-422X-9-235.

Tamura, K., Peterson, D., Peterson, N., Stecher, G., Nei, M., and Kumar, S. 2011. MEGA5: Molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods. Mol. Biol. Evol. 28:2731–2739. doi:10.1093/molbev/msr121.

Thekke-Veetil, T., Ghanem-Sabanadzovic, N. A., Keller, K. E., Martin, R. R., Sabanadzovic, S., and Tzanetakis, I. E. 2013. Molecular characterization and population structure of blackberry vein banding associated virus, new ampelovirus associated with yellow vein disease. Virus Res. 178:234-240. doi:10.1016/j.virusres.2013.09.039.

Tatineni, S., Ziems, A. D., Wegulo, S. N., and French, R. 2009a. Triticum mosaic virus: A distinct member of the family Potyviridae with an unusually long leader sequence. Phytopathol. 99:943-950. doi:10.1094/PHYTO-99-8-0943.

Tatineni, S., Afunian, M. R., Gowda, S., Hilf, M. E., Bar-Joseph, M., and Dawson, W. O. 2009b. Characterization of the 5′- and 3′-terminal subgenomic RNAs produced by a capillovirus: Evidence for a CP subgenomic RNA.Virology 385:521-528.

Thompson, J. D., Higgins, D. G., and Gibson, T. J. 1994. CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position- specific gap penalties and weight matrix choice. Nucleic Acids Res. 22:4673–4680. doi:10.1093/nar/22.22.4673.

Thompson, J. R., Fuchs, M., and Perry, K. L. 2012. Genomic analysis of grapevine leafroll associated virus-5 and related viruses. Virus Res. 163:19–27. doi:10.1016/j.virusres.2011.08.006.

Tian, T., Rubio, L., Yeh, H. H., Crawford, B., and Falk, B. W. 1999. Lettuce infectious yellows virus: invitro acquisition analysis using partially purified virions and the whitefly Bemisia tabaci. J. Gen. Virol. 80:1111-1117. doi:10.1099/0022-1317-80-5-1111.

Ullmann, A., Jacob, F., and Monod, J. 1967. Characterization by in vitro complementation of a peptide corresponding to an operator-proximal segment of the beta-galactosidase structural gene of Escherichia coli. J. Mol. Biol. 24:339–343. doi:10.1016/0022-2836(67)90341-5.

Valverde, R. A, Nameth, S. T., and Jordan, L. R. 1990. Special topic analysis of double-stranded RNA for plant virus diagnosis. Plant Dis. 74:255–258.

Vieira, J., and Messing, J. 1982. The pUC plasmids, an M13mp7-derived system for insertion mutagenesis and sequencing with synthetic universal primers. Gene 19:259–268.

Vitushkina, M. V., Rogozin, I. B., Jelkmann, W., Koonin, E. V., and Agranovsky, A. A. 2007. Completion of the mapping of transcription start sites for the five-gene block subgenomic

85

RNAs of Beet yellows Closterovirus and identification of putative subgenomic promoters. Virus Res. 128:153–158.

Velasco, L., Cretazzo, E., Padilla, C. V., and Janssen, D. 2015. Grapevine leafroll associated virus 4 strain 9: Complete genome and quantitaive analysis of virus-derived small interfering RNA populations. Journal of Plant Pathology 97: 189-192. doi:10.4454/JPP.V97I1.051. van den Born, E., Omelchenko, M. V., Bekkelund, A., Leihne, V., Koonin, E. V., Dolja, V. V., and Falnes, P. O. 2008. Viral AlkB proteins repair DNA damage by oxidative demethylation. Nucleic Acids Res 36: 5451-5461. doi:10.1093/nar/gkn519.

Warren, C. E., and Murphy, J. F. 2003. The complete nucleotide sequence of Pepper mottle virus-Florida RNA. Arch. Virol. 148:189-97. doi:10.1007/s00705-002-0915-2.

Yeh, H. -H., Tian, T., Rubio, L., Crawford, B., and Falk, B. W. 2000. Asynchronous accumulation of Lettuce Infectious Yellows Virus RNAs 1 and 2 and identification of an RNA1 trans enhancer of RNA 2 accumulation. J. Virol. 74:5762–5768.

Zhu, H. Y., Ling, K. S., Goszczynski, D. E., McFerson, J. R., and Gonsalves, D. 1998. Nucleotide sequence and genome organization of grapevine leafroll-associated virus-2 are similar to beet yellows virus, the closterovirus type member. J. Gen. Virol. 79:1289–1298. doi:10.1099/0022-1317-79-5-1289.

Zuker, M., Mathews, D. H., and Turner, D. H. Algorithms and thermodynamics for RNA secondary structure prediction. A practical guide. 1999. RNA Biochemistry and Biotechnology. NATO Science Series. pp.11-43.

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CHAPTER THREE

SPATIO-TEMPORAL SPREAD OF GRAPEVINE LEAFROLL-ASSOCIATED VIRUS 3

IN WASHINGTON VINEYARDS

Bhanu Priya Donda, Sandya Kesoju, Neil Mc Roberts and Rayapati A. Naidu

(Unpublished)

ABSTRACT

The spread of grapevine leafroll disease (GLD) to vineyards planted with certified planting stock is of significant concern to the grape and wine industry in Washington State. In this study, the spatial and temporal spread of GLD was examined in three young vineyard blocks planted with virus-tested wine grape cultivars in two geographic locations adjacent to vineyard blocks heavily infected with GLD. During each season, the position of vines showing GLD symptoms was recorded in a matrix representing the planting lattice. Symptomatic vines were tested positive only for Grapevine leafroll-associated virus 3 (GLRaV-3), the ubiquitous virus in

Washington vineyards. The results from several seasons showed a gradual increase in disease incidence over successive seasons in all three young blocks. Spatial and temporal mapping of

GLD indicated a disease gradient in which the highest percentage of symptomatic vines were in rows proximal to infected old blocks. Spatial autocorrelation (dependence) analysis suggested random distribution of symptomatic vines in the three blocks during initial years indicating primary spread of the virus and clustering of symptomatic vines during subsequent years, suggesting vine-to-vine secondary spread within the block. Molecular analysis of GLRaV-3 variants revealed the spread of distinct genetic variants of the virus into new plantings. These

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results provided insights on the spatial and temporal spread of GLD to facilitate designing site- specific disease management strategies.

INTRODUCTION

Grapevine leafroll disease (GLD) has arguably received more attention than other virus diseases affecting grapevines (Vitis spp.) due to its ubiquitous distribution in grapevine-growing regions worldwide (Martelli, 2014; Naidu et al., 2014). The disease affects a wide range of V. vinifera cultivars, with red-berried cultivars showing conspicuous symptoms compared to white- berried cultivars (Rayapati et al., 2008). The disease is known to induce a wide range of disorders, vine growth problems, reduced yield, delayed fruit maturity and poor quality of grapes

(Goheen, 1970; Bovey et al., 1980; Bovey and Martelli, 1992; Golino et al., 2009 a & b; Lee and

Martin, 2009; Lee et al., 2009; Basso et al., 2010; Komar et al., 2010; Alabi et al., 2012) resulting in significant financial losses to grower’s income (Atallah et al., 2012; Ricketts et al.,

2015). Within Washington State, a recent economic study estimated that a commercial wine grape grower could lose up to $20,000 per acre over the 20-year period depending on the magnitude of annual reduction in fruit yield and decline in fruit quality (Naidu and Walsh, 2015).

Several morphologically similar but genetically distinct viruses, designated as grapevine leafroll-associated viruses (GLRaVs), have been documented in grapevines exhibiting GLD symptoms or suspected for the disease (Martelli et al., 2002; Martelli, 2014; Naidu et al., 2014).

GLRaVs are phloem-limited belonging to the family Closteroviridae, with GLRaV-1, -3, and -4 assigned to the genus Ampelovirus, GLRaV-2 to the genus Closterovirus, and GLRaV-7 to the genus Velarivirus (Al Rwahnih et al., 2012; Martelli et al., 2012; Martelli, 2014; Naidu et al.,

2015). Among them, GLRaV-3 is considered the most ubiquitous virus species in all grape- producing regions around the world (Maree et al., 2013). Likewise, GLRaV-3 was found to be

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the most predominant among the four GLRaV species documented in Washington vineyards

(Naidu, 2011).

GLRaVs are mainly disseminated over long distances via distribution of infected planting stock. In addition, GLRaV-1, -3 and -4 and its strains are known to be spread by phloem-feeding mealybugs (Hemiptera: Pseudococcidae) and soft scales (Hemiptera: Coccidae) in a semi- persistent manner (Tsai et al., 2010; Almeida et al., 2013; Naidu et al., 2014 and cited references). Although several species of mealybugs were implicated as vectors of these ampeloviruses (Naidu et al., 2014 and cited references), only grape mealybug (Pseudococcus maritimus) (Hemiptera: Pseudococcidae) was reported to be present in Washington State vineyards (Bahder et al., 2013a). Recently, the grape mealybug (P. maritimus) and European fruit lecanium scale (Parthenolecanium corni) were reported to be capable of transmitting

GLRaV-3 in Washington State vineyards (Bahder et al., 2013b). In contrast, no insect vectors have so far been reported for GLRaV-2 and -7, although the former can be transmitted by manual inoculations to Nicotiana benthamiana (Goszczynski et al., 1996) and the latter via plant parasitic cuscuta species (Mikona and Jelkmann, 2010). Although direct planting of infected cuttings or grafting of infected materials (either rootstock or scion or both) can lead to the introduction of all GLRaVs into newly planted vineyards, only ampeloviruses are transmitted by mealybugs and soft scales contributing to their natural spread between and within vineyards

(Naidu et al., 2014). Among the ampeloviruses, GLRaV-3 appears to play a significant role in the epidemiology of GLD, likely due to its efficient transmission by several species of mealybugs and scale insects (Golino et al., 2002; Mahfoudhi et al., 2009; Tsai et al., 2010;

Almeida et al., 2013; Naidu et al., 2014 and cited references). GLRaV-3 is known to occur as distinct variant groups, some of which can cause serious symptoms and others causing

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asymptomatic infections (Fajardo et al., 2007; Fuchs et al., 2009; Jooste et al., 2011; Gouveia et al., 2011; Sharma et al., 2011; Bester et al., 2012; Farooq et al., 2013; Liu et al., 2013; Maree et al., 2015). Due to its dominant and worldwide distribution, GLRaV-3 received greater attention and considered as the most economically important virus than other GLRaVs for grape production.

Due to the lack of curative measures, preventive measures are commonly applied for minimizing the spread of GLRaVs (Pietersen et al., 2013). The use of virus-tested planting stock is recommended as the first line of defense in preventing the introduction of GLRaVs into newly planted vineyard blocks. Although this is effective in preventing the dissemination of all

GLRaVs via propagation materials, controlling insect vectors have been advocated for post- planting management of GLD to reduce the spread of ampeloviruses by mealybugs and soft scales (Almeida et al., 2013; Naidu et al., 2014). However, recent studies have shown that application of insecticides have provided partially successful vector control measures to slow the spread of GLD (Daane et al., 2012; Wallingford et al., 2015). Besides, roguing or removing infected vines and replacing with cuttings derived from certified virus-tested mother vines has been commonly advocated to eliminate sources of inoculum for reducing the spread of GLD within vineyards.

Studies conducted in several grapevine-growing regions have made important advances in understanding various aspects of the epidemiology of GLD (Engelbrecht and Kasdorf, 1985;

Habili and Nutter, 1997; Charles et al., 2009; Gribaudo et al., 2009; Sokolsky et al., 2013; reviewed in Almeida et al., 2013). Due to the wide spread occurrence of GLRaV-3 and its significant contributing role to GLD symptoms, all these studies were focused mainly on

GLRaV-3. Available information indicates that the spread of GLRaV-3 occurs via two main

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pathways (Naidu et al., 2014). Primary spread can occur via virus-infected planting stock or initiated by viruliferous vectors moving on their own, dispersed by wind and/or carried on vineyard implements and workers’ clothes. This could result in random distribution of infected vines in the vineyard block. Conversely, secondary spread within a vineyard can occur due to vecor transmission of GLRaV-3 leading to an increased number of symptomatic vines over time and space. The pattern of spatial distribution of GLD within an infected vineyard can show clustering of infected vines suggestive of vine-to-vine secondary spread along the rows or ‘edge effect’ where a disease gradient occurs with high percentage of symptomatic vines located at the edges of a vineyard and the disease incidence tapering towards the middle or the other side of the vineyard (Charles et al., 2006; Sokolsky et al., 2013). However, no information is currently available on the spread of GLD in Washington vineyards.

In this study, multi-season observations were made to analyze spatial patterns of GLD spread within new vineyard blocks planted in proximity to old blocks that are heavily infected with the disease. In addition, genetic diversity of GLRaV-3 was examined to evaluate the significance of strain variation in the context of GLD spread within new plantings. The results provided insights on the spatial and temporal spread of GLD to facilitate designing site-specific disease management strategies in vineyards.

MATERIALS AND METHODS

Study sites and wine grape cultivars

Three commercial vineyard blocks, each planted with own-rooted V. vinifera cvs.

Cabernet Sauvignon, Syrah and Petit Syrah, were selected for this study. The Cabernet

Sauvignon and Syrah blocks were in one location and the Petit Syrah block was in a geographically separate location. Specific approval was obtained from the vineyard owners for

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collecting the data used this study. Vineyard locations and names of property owners were withheld due to grower confidentiality.

The 1.416 hectare (14,162 m2) Cabernet Sauvignon block was planted in 2007 and contains 26 rows with each row having about 100 vines. The planting matrix was 1.83 m between neighboring vines along the rows and 2.74 m between adjacent rows. The block was surrounded on North and East with approximately 30 year old Chardonnay and 50 year old

Cabernet Sauvignon blocks heavily infected with GLD and on West and South with Merlot and

Semillon blocks planted with virus-tested planting stock in 2007 (Fig. 14A). The young Cabernet

Sauvignon block was separated from the neighboring Cabernet Sauvignon, Chardonnay and

Semillon blocks by 12.19 m dirt road, whereas the Merlot block is an extension of the young

Cabernet Sauvignon block with 2.74 m between them.

The 1.145 hectare (11,451 m2) Syrah block was planted in 2004 and contains 28 rows with each row having about 95 vines. The planting matrix was 3.05 m between rows and 1.83 m between adjacent vines within rows. The block was surrounded on South and West with approximately 15 and 30 years old Cabernet Sauvignon blocks and on North with approximately

25 year old Merlot block (Fig. 15A). All these blocks were heavily infected with GLD. An apple orchard was present on the East side of the block. The Syrah block was separated from old

Cabernet Sauvignon on West side by 9.14 m dirt road and Merlot block on North side by 12.80 m dirt road, whereas the Cabernet Sauvignon on South side is an extension of the young Syrah block with 3.05 m distance between them.

The 0.785 hectare (7,849 m2) Petit Syrah block was planted in 2009 and contains 17 rows with each row having about 92 vines. The planting matrix was same as in Syrah block mentioned above. This block was surrounded with an approximately 30 year old Cabernet Sauvignon block

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on West and an approximately 25 year old Chardonnay block on South with both blocks heavily infected with GLD (Fig. 16A). The Chardonnay block was separated by a 10.67 m dirt road. A young Zinfandel block planted in 2009 is located on the East separated by a distance of 3.05 m, whereas the North side of the Petit Syrah block is an open field.

Spatial and temporal mapping of symptomatic vines

Cabernet Sauvignon, Syrah and Petit Syrah blocks were monitored for GLD symptoms annually between the last week of September and first week of October, when symptom expression was optimal in these cultivars under conditions prevailing in Washington State.

Cabernet Sauvignon was monitored between 2007 and 2015, Syrah block between 2008 and

2015 and Petit Syrah between 2012 and 2015. The Syrah block was not monitored in 2011 due to inclement weather that severely damaged canopy in this block. During each season, the three vineyard blocks were monitored for GLD symptoms and the position of individual vines showing symptoms were recorded and plotted in a XY matrix representing the planting lattice using the row number and vine position as coordinates. The three blocks were georeferenced with the global positioning system (GPS). Using the distance between rows and between vines within each row, the spatial distribution of all symptomatic and nonsyptomatic vines in each block was mapped using ArcGIS ver. 10.2 software (Esri, Redlands, California). Annual maps specific to each block were prepared for each season using cumulative data of new infections from that season and the preceding seasons.

Statistical analysis of the spatio-temporal data:

Temporal analysis

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Number of symptomatic vines recorded during each season in all three vineyard blocks was used to calculate the proportion of infected vines using the following equation:

Proportion of infection = No/Nt

Where,

No = total number of symptomatic vines in a given season and

Nt = total number of vines in a vineyard block

Disease progress curves and regression analyses were used to determine the most appropriate model for quantifying temporal disease spread within vineyard blocks. Disease progress curves were subjected to the exponential, monomolecular, logistic, and Gompertz models to determine a suitable model that would best fit the relationship between GLD incidence and time (SAS version 6.04; SAS Institute, Inc., Cary, North Carolina; Nutter and Parker, 1997;

Habili and Nutter, 1997). The best fit model was selected based on the Akaike information criterion (AIC), Bayesian information criterion (BIC), and root mean squared error (RMSE).

Based on initial analyses, the Gompertz model was judged to be the most appropriate for describing disease spread with time in all the three vineyard blocks. However, the logistic model was also found to give acceptable data sets for analyzing disease spread over time. Temporal analyses of the data set from individual blocks were carried out using JMP 7.0 (SAS Institute,

Cary, North Carolina).

Spatial analysis

Spatial autocorrelation (dependence) analysis was performed using number of infected vines in a given spatial grid (3x3 m) to test whether symptomatic vines were distributed randomly or clustered across the block with ArcMap ver. 10.2 software (ESRI Redlands,

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California). Subsequently, the data was subjected to statistical significance (Kulldorff, 1997).

The Moran’s I Index (Moran’s I) statistics (Boots and Getis, 1998) was used to evaluate significant clustering (p < 0.05) of infected vines within the block. The value of Moran’s I range from −1 to +1 with positive Moran’s I values above zero considered as clustering (aggregation) and a negative value as dispersed distribution, while zero value taken as random distribution.

Sample collection, reverse transcription PCR (RT-PCR), cloning and sequencing for

GLRaV-3

Initially, leaf samples were collected from symptomatic and adjacent nonsymptomatic vines in Cabernet Sauvignon, Syrah and Petit Syrah blocks and tested for the presence of

GLRaV-1, -3 and -4 by one step reverse transcription-polymerase chain reaction (RT-PCR) assay using virus-specific diagnostic primers (Bahder et al., 2013a). The results indicated the presence of only GLRaV-3 in samples collected from symptomatic vines and absence of all

GLRaVs in nonsymptomatic vines (data not shown). Due to the tight correlation between symptoms and the presence of GLRaV-3, only symptomatic samples from the three blocks were collected each season for testing by RT-PCR for the presence of GLRaV-3. In addition, samples were collected from symptomatic vines in the three blocks as well as in the neighboring heavily infected old blocks for analyzing sequence variation in the heat-shock protein 70 homolog

(HSP70h) gene encoded by GLRaV-3 (Jarugula et al., 2010). For this purpose, gene-specific primers were designed based on GLRaV-3 sequences available in GenBank (GLRaV-3 variant group I: NC_004667; group II: EU259806; group III: GQ352633, JQ423939; group VI:

JQ655295, JQ655296, JX220899, JX220900; group VII: KM0589745). The forward primer (5’-

GGGGDGGRACTTTCGAYGTSTC-3’) corresponds to 11260 - 11281 nt and the reverse primer

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(5’-ATTGGACTRCCYTTYGGGAAAAT-3’) is complementary to 11844-11866 nt in the genome of GLRaV-3 isolate WA-MR (GU983863, Jarugula et al., 2010). In one step RT-PCR assays (Bahder et al., 2013a), these primers amplified approximately 600 bp DNA fragment specific to the HSP70h gene. The PCR amplified virus-specific fragments were cloned into

TOPO 2.1 vector (ThermoFischer Scientific, Grand Island, New York) and transformed into

Escherichia coli. Plasmid DNA from positive recombinant colonies was purified using the

QIAprep spin miniprep kit (Qiagen Inc., Valencia, California). Three independent clones per sample were sequenced in both orientations and a consensus sequence was generated when three independent clones showed ≥98% identities. Additional clones were sequenced from samples that showed ≤98% nucleotide sequence identity between the three initially sequenced clones to analyze possible occurrence of mixed infection of different variants of GLRaV-3. The sequences were aligned along with the reference sequences for different variant groups extracted from

GenBank (GLRaV-3 variant group I: NC_004667; group II: EU259806; group III: GQ352633,

JQ423939; group VI: JQ655295, JQ655296, JX220899, JX220900; group VII: KM0589745) using CLUSTAL W multiple sequence alignment (Thompson et al., 1994). The sequence of

GLRaV-3 variant group VIII, reported in a recent publication by Maree et al., 2015, is not accessible in GenBank and, therefore, not included in this analysis. The sequence corresponding to HSP70h of GLRaV-1 (Accession number NC_016509) was used as an out group. Aligned sequences were used to build a neighbor-joining phylogenetic tree with MEGA5 software

(Tamura et al., 2011). A total of 1,000 boot strap replications were used to assess robustness of the phylogenetic tree.

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RESULTS

Correlation between GLD symptoms and the presence of GLRaV-3

The characteristic symptoms of GLD were observed in all three cultivars monitored during each season. These symptoms matched with those reported earlier (Rayapati et al., 2008) and made it relatively easy to identify symptomatic vines by visual observations during each season. However, symptom-based diagnosis is complicated by the fact that at least four species of GLRaVs (GLRaV-1, -2, -3 and -4) have so far been reported in grapevines showing GLD symptoms in Washington vineyards (Naidu, 2011). Thus, in order to fully characterize GLRaVs present in GLD-infected grapevines, representative samples were collected from both symptomatic and nonsymptomatic vines during initial stages of this study and tested by RT-PCR for the presence of GLRaV-1, -2, -3 and -4. The results indicated the presence of only GLRaV-3 in symptomatic, but not in nonsymptomatic vines in Cabernet Sauvignon, Syrah and Petit Syrah blocks (Fig. 17). These results provided good correlation between GLD symptoms and the presence of GLRaV-3 and further supported previous observations that GLRaV-3 is the primary virus species associated with GLD in Washington vineyards (Naidu, 2011). Based on these results, diseased grapevines were identified during each season based on visual symptoms.

Spatial and temporal patterns of GLD spread into and within vineyards:

Temporal analysis

A gradual increase in disease incidence in terms of increased number of symptomatic vines in an otherwise healthy vineyard block over successive seasons could be due to new infections from external sources. Thus, monitoring GLD incidence in newly planted healthy vineyards and analyzing their spatial pattern should provide valuable epidemiological

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information to infer sources of inoculum and analyze the influence of various biotic and climatic factors in the spread GLD across Washington vineyards. Towards this objective, vineyard blocks planted with Cabernet Sauvignon, Syrah and Petit Syrah blocks were monitored for the spread of

GLD during consecutive seasons between 2007 and 2015.

In Cabernet Sauvignon block, a few symptomatic vines positive for GLRaV-3 were observed between 2008 and 2011 within the first five rows on the East side of the block, followed by a gradual increase in number of symptomatic vines within these rows during 2012 and 2015 (Fig. 14B). In addition, new infections were observed in other rows throughout the block between 2012 and 2015. The spatial map depicting cumulative data at the end of 2015 showed higher number of symptomatic vines located on the East side followed by North side indicating that GLD spread likely occurred from heavily infected old Cabernet Sauvignon and

Chardonnay block located, respectively, on the East and North side with respect to the young

Cabernet Sauvignon block. GLD incidence during the eight seasons showed slow progression in the beginning (2008 to 2010) followed by increased spread during the subsequent years (2011 to

2015) (Fig. 18). Though the cumulative disease incidence in the block was about 0.09 (9%) at the end of 2015 (Table 8A), the Gompertz model predicted that the maximum spread of GLD could continue and reach up to about 0.28 (28%) at the end of 2016 (Table 9). These results suggest that the disease progress is in its exponential phase and could transition to the lag phase in future giving the sigmoid shape of the disease curve.

Although the Syrah block was not monitored from the beginning of planting, observations made from 2008 showed the presence of higher number of symptomatic vines positive for GLRaV-3 within the first 2-3 rows on the South that are adjacent to an old Cabernet

Sauvignon block (Fig.15B). In subsequent seasons, increased number of symptomatic vines was

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observed on the South side of the block followed by North and West sides. This spatial variation in the number of symptomatic vines across the block could be due to the distance between infected old blocks and the young Syrah block. Since the heavily infected old Cabernet

Sauvignon block on the South side is located more closely than the other two old blocks on the

North and West side, respectively, it can be concluded that close proximity to an infected block increases the likelihood of virus spread into an apparently healthy vineyard. The Gompertz model showed a gradual increase in the disease incidence over the seven year period in Syrah block (Fig. 19). The model predicted that the maximum spread could reach 100 percent

(predicted value of 1.21 or 121%) at the end of 2019 (Table 10), though the cumulative disease incidence in the block was about 0.25 (25%) at the end of 2015 (Table 8B). Similar to the young

Cabernet Sauvignon block, the disease progress is in its exponential phase and could transition to the lag phase in future giving the sigmoid shape of the disease curve.

The Petit Syrah block planted in 2009 was monitored between 2012 and 2015. Like in the other two blocks described above, the spatial distribution of GLD showed higher number symptomatic vines located in rows close to the heavily infected old blocks than in distal parts of the Petit Syrah block. The overall data on temporal spread of GLD in this block (Fig. 20) followed a trend similar to that observed in Cabernet Sauvignon and Syrah blocks (Table 8C).

However, the cumulative incidence of GLD at the end of 2015 was 0.42 (42%) in this block, which is far greater than the cumulative incidence in the other two blocks.

Gompertz model was used to analyze the temporal disease progress in the three vineyard blocks. The S-shaped sigmoidal growth curve, consisting of an initial slow infection rate followed by exponential/logarithmic growth phase transitioning to a final linear/saturation phase, has been used previously in studying the temporal spread of GLD (Jordan, 1993; Habili and

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Nutter, 1997; Sokolsky et al., 2013). In Petit Syrah block, the Gompertz model estimated maximum spread at 1.99 (199%) at the end of 2017 (Table 11). The parameter estimates of

Gompertz model for all the three blocks predicted maximum spread after 2015, indicating that

GLD spread could continue at higher rate after 2015. In Cabernet Sauvignon block the disease spread increased until the inflection point is reached i.e., until 2016; Syrah block until 2019; and

Petit Syrah until 2017 (Tables 9, 10, 11).

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A

B

Figure 14. Spatial and temporal spread of GLD in a young Cabernet Sauvignon block. (A)

Google map showing the newly planted Cabernet Sauvignon block (yellow rectangle border) surrounded by heavily GLD-infected old blocks (red rectangle borders). (B) The spatial and temporal distribution of GLD during 2008 and 2015. The red circles represent symptomatic vines and the green circles indicate nonsymptomatic vines.

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B

A

Figure 15. Spatial and temporal distribution of GLD in a young Syrah block. (A) Google map showing the young Syrah block (yellow rectangle border) and heavily infected old blocks (red rectangle border). (B) The spatio-temporal distribution of GLD during 2008 and 2015. The red circles represent GLD symptomatic vines and the green circles indicate nonsymptomatic vines.

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B

A

Figure 16. Spatial and temporal distribution of GLD in a young Petit Syrah block. (A) Google map showing the young Petit Syrah block (yellow rectangle border) and heavily infected old blocks (red rectangle border). (B) The spatio-temporal distribution of GLD during 2012 and

2015. The red circles represent GLD symptomatic vines and the green circles indicate nonsymptomatic vines.

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A

B

Figure 17. Correlation between GLD symptoms and the presence of GLRaV-3. (A) Genome map of GLRaV-3 showing the HSP70h region targeted for RT-PCR amplification. The genome map and location of open reading frames encoded by the virus are described in Jarugula et al. (2010).

(B) An agarose gel showing an approximately 600 base pair DNA band (shown by arrow on the left) amplified from symptomatic (Lane 1-4) but not from non-symptomatic vines (Lane 5-8). Mr

= 1 kb plus DNA molecular weight markers to estimate the size of amplified DNA fragment. ‘+’ and ‘–’ represent positive and negative controls, respectively.

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Table 8A:

Total number of Total number of vines Proportion of Year symptomatic vines (No) (Nt) infection* 2008 4 2592 0.0015 2009 10 2592 0.0039 2010 17 2592 0.0066 2011 30 2592 0.0116 2012 81 2592 0.0313 2013 127 2592 0.0490 2014 146 2592 0.0563 2015 226 2592 0.0872

Table 8B:

Total number of GLD Total number of vines Proportion of Year symptomatic vines (No) (Nt) infection* 2008 46 2090 0.0220 2009 83 2090 0.0397 2010 120 2090 0.0574 2012 232 2090 0.1110 2013 280 2090 0.1339 2014 378 2090 0.1809 2015 532 2090 0.2545

Table 8C:

Total number of GLD Total number of vines Proportion of Year symptomatic vines (No) (Nt) infection* 2012 69 1564 0.0441 2013 237 1564 0.1515 2014 380 1564 0.2430 2015 659 1564 0.4214

Table 8. Annual incidence of GLD in the three vineyard blocks. Cumulative number of symptomatic vines recorded each year, total number of vines in the block and proportion of infection are shown for Cabernet Sauvignon (A), Syrah (B) and Petit Syrah (C) blocks.

* See materials and methods section for the mathematical equation.

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Figure 18. Temporal increase in GLD incidence in Cabernet Sauvignon block during 2008 and

2015 seasons.

Parameter Estimate Standard error Lower 95% Upper 95% Asymptote 0.284 0.260 -0.225 0.783 Growth rate 0.216 0.110 0.001 0.432 Inflection point 2016 3.835 2008 2023

Table 9. Parameter estimates for disease incidence in the Cabernet Sauvignon block using

Gompertz model.

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Figure 19. Temporal increase in GLD incidence in the Syrah block during 2008 and 2015 seasons.

Parameter Estimate Standard error Lower 95% Upper 95% Asymptote 1.212 1.504 -1.735 4.160 Growth rate 0.133 0.074 -0.013 0.279 Inflection point 2019 7.623 2003 2033

Table 10. Parameter estimates for disease incidence in the Syrah block using Gompertz model.

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Figure 20. Temporal increase in GLD incidence in the Petit Syrah block during 2012 and 2015

Parameter Estimate Standard error Lower 95% Upper 95% Asymptote 1.992 4.109 -6.061 10.046 Growth rate 0.275 0.272 -0.258 0.808 Inflection point 2017 6.343 2004 2029

Table 11. Parameter estimates for disease incidence in the Petit Syrah block using Gompertz model.

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Spatial analyses

To determine whether the patterns of GLD spread in Cabernet Sauvignon, Syrah and Petit

Syrah blocks were random and/or aggregated over time, the spatial data (Figs. 14, 15 and 16) was analyzed using spatial autocorrelation. Disease incidence in Cabernet Sauvignon during

2008 and 2009 was not modeled due to lower number of symptomatic vines. In Cabernet

Sauvignon and Syrah blocks, significant positive Moran’s I values were observed from 2012 and

2010, respectively, indicating clustering of symptomatic vines from these time periods in the two blocks (Tables 12 and 13). Similar analysis was not carried out with Petit Syrah block due to the data available for a limited number of years. These results indicated random distribution of symptomatic vines during initial growing seasons (until 2011 and 2009 in Cabernet Sauvignon and Syrah blocks, respectively), but showed significant autocorrelation or aggregated spatial pattern in subsequent seasons beginning from 2012 in Cabernet Sauvignon and 2010 in Syrah blocks. Based on the spatial autocorrelation analyses presented here, it can be concluded that random distribution of symptomatic vines in young blocks during initial years is an indication of primary spread of GLD, whereas aggregation or clustering within and across rows during subsequent years could be due to vine-to-vine secondary spread of the disease within a vineyard.

Distribution of GLRaV-3 variants in young vineyard blocks

Like other RNA viruses, genetic variants of GLRaV-3 have been reported in several grapevine-growing regions (Turturo et al., 2005; Fajardo et al., 2007; Engel et al., 2008; Fuchs et al., 2009; Gouveia et al., 2011; Sharma et al., 2011; Bester et al., 2012; Chooi et al., 2013). The available information indicates the presence of three to eight variant groups, identified based on the HSP70 and CP gene sequences in many regions around the world (Turturo et al., 2005;

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Fajardo et al., 2007; Fuchs et al., 2009; Gouveia et al., 2011; Jooste et al., 2011; Sharma et al.,

2011; Wang et al., 2011; Bester et al., 2012; Chooi et al., 2013; Farooq et al., 2013; Liu et al.,

2013; Maree et al., 2015). However, information on the occurrence and spread of GLRaV-3 variants in Washington vineyards is not available. In this study, therefore, the presence of

GLRaV-3 variants was examined to determine whether a single variant group is predominantly spreading into new blocks. For this purpose, a total of 46, 56 and 66 samples were collected, respectively, from Cabernet Sauvignon, Syrah and Petit Syrah blocks, representing 10 to 20% of the total symptomatic vines in the three blocks. Samples were collected randomly from the symptomatic vines in each vineyard block between 2012 and 2015 and used to analyze the partial sequence of the HSP70h gene. In addition, HSP70h gene sequences derived from a total of 190 samples collected from adjacent old blocks were included in this study. The results presented in Figs. 21 and 22 indicated that sequences belonging to variant group I were found in

~58%, ~55% and ~85% of the samples analyzed from Cabernet Sauvignon, Syrah and Petit

Syrah blocks, respectively. Similarly, virus sequences in adjacent old blocks were found belonging to group I. Samples that were not grouped with the reference sequences in the phylogenetic tree constructed based on HSP70h region of GLRaV-3 (Maree et al., 2015) were shown as not grouped (NG). Mixed infections of group I with other variant groups, such as group

II, III, VI and NG, were observed in low number of samples. Group VII was not found in samples analyzed from the three vineyard blocks. Altogether, these results indicated that group I isolates are predominant in the three blocks and adjacent old blocks compared to other variant groups of GLRaV-3.

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Year Moran's I z-score p-value Pattern 2010 -0.050113 0.235986 0.813444 Random 2011 -0.016154 0.183836 0.854142 Random 2012 0.244854 4.173711 0.00003 Clustered 2013 0.065589 2.006221 0.044833 Clustered 2014 0.095306 2.667543 0.007641 Clustered 2015 0.058568 2.250998 0.024386 Clustered

Table 12. Spatial autocorrelation (dependence) analysis using Moran’s I index for GLD incidence in the Cabernet Sauvignon block. Significant positive spatial autocorrelation occurs at a particular distance when P ≤ 0.05. Dispersed or random pattern occurs when p > 0.05.

Year Moran's I z-score p-value Pattern 2008 -0.156154 -0.450969 0.652012 Random 2009 -0.006515 0.203241 0.838947 Random 2010 0.111135 3.904411 0.000094 Clustered 2012 0.123313 3.484092 0.000494 Clustered 2013 0.166868 4.876135 0.000001 Clustered 2014 0.127486 4.084388 0.000044 Clustered 2015 0.077755 2.933235 0.003354 Clustered

Table 13. Spatial autocorrelation (dependence) analysis using Moran’s I index for GLD incidence in the Syrah block. Significant positive spatial autocorrelation occurs at a particular distance when P ≤ 0.05. Dispersed or random pattern occurs when p > 0.05.

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A

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B

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C

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Figure 21. Phylogenetic analysis of GLRaV-3 sequences based on partial nucleotide sequences of the HSP70h gene. The Neighbor-

joining phylogenetic trees were constructed with GLRaV-3 sequences obtained from Cabernet Sauvignon (A), Syrah (B) and Petit

Syrah (C) blocks and neighboring heavily infected old blocks. The reference sequences for GLRaV-3 groups I, II, III and VI in the

phylogram are shown in bold. NG= Not Grouped. The trees were rooted using corresponding sequence of GLRaV-1. Boot strap values

(1,000 replicates) are shown at the branch nodes.

11 5

Figure 22. Pie diagram showing the proportion of GLRaV-3 variants analyzed from samples collected from the three vineyard blocks

(cvs. Cabernet Sauvignon, Syrah and Petit Syrah) and adjacent old blocks. Variant groups I-VI are as described in Maree et al., 2015.

NG = variants that did not belong to documented variants groups.

DISCUSSION

This study provides for the first time the spatial and temporal spread of GLD in young vineyards under conditions prevailing in Washington State. The temporal progress of GLD in three young vineyard blocks followed similar trends in that vineyard blocks planted in close proximity to heavily infected old blocks had a higher probability of being infected with GLD

(Figs. 14, 15 and 16). In addition, the spatial dynamics of disease spread was mostly random during the initial stages following by exponential increase leading to aggregation or clustering of infected vines. However, the results indicated that the proportion of infected vines in each block and the rate at which new infections occur during each season varied widely among the three cultivars. These results also indicated variability in disease incidence across geographic locations and between seasons. One explanation for this difference could be the sources of virus inoculum surrounding the new plantings. It is likely that higher incidence in the Syrah block (Fig. 19) could be due to the presence of heavily infected old blocks on three sides. However, this cannot account for higher incidence in Petit Syrah block (Fig. 20), where rapid increase in disease incidence within a short period of time could be due to other confounding factors, such as wind and other site-specific factors.

The spatio-temporal distribution suggested a disease gradient where the disease incidence decreased with increase in the distance from the old block. This gradient of infection originating from the adjacent old blocks to the margins of young vineyard blocks emphasizes the spread of

GLD due to immigrating first instar stages of mealybugs either by their own motility over short distances, or on farm workers clothing, on implements, by wind, or possibly even by birds

(Pietersen, 2006; Almeida et al., 2013). Furthermore, an increase in clustering was observed each year in the three vineyard blocks. Such slow but constant plant to plant transmission is indicative

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of secondary spread due to slow moving vectors such as mealybugs and scale insects (Cabaleiro et al., 2008). Previous studies have suggested that the secondary spread from an infected vine to a healthy vine is similar when the neighboring vine is within or across a row from the infected vine (Sokolsky, 2013). This secondary spread is further dependent on the active or passive movement of the vectors and its efficiency in transmitting the virus. In case of mealybugs, the first instar stages are reported to be more efficient in transmitting GLRaV-3 (Tsai et al., 2008).

The grape mealybug (Ps maritimus) found in the Washington vineyards has only 3 instar larval stages as opposed to 9 instar stages in vine mealybug (Planococcus ficus), which was first introduced in Southern California in early 1900’s and became a major pest in the later years

(Daane et al., 2004). Thus, the temporal dynamics of the disease spread reported in this study might be different from other locations, depending on the vector species and grapevine cultivar.

GLD is a complex disease with varying number of GLRaVs, its strains and variants co- existing in a single grapevine. This affects the disease spread by the vectors. Previous reports have suggested that GLRaV-3 was the most predominant and widely distributed virus among the

GLRaVs present in WA vineyards (Naidu, 2011). Furthermore, the samples tested from symptomatic and non-symptomatic vines also indicated a good correlation between GLD symptoms and the presence of GLRaV-3. Therefore, identifying GLRaV-3 variants in the samples from old and young vineyard blocks was the main focus of this study. Since it is difficult to sequence every sample from an infected vine in young and adjacent old vineyard blocks, this study was limited to a select number of symptomatic samples (10 to 20 %) for analyzing sequence variation in the HSP70h gene of GLRaV-3. Previous study showing the genetic diversity of GLRaV-3 in South African vineyards has indicated that variant group II was predominantly found in the vineyard clusters studied from the mother blocks (Jooste et al.,

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2011). In Napa Valley region in California, group I (27% of isolates studied) and III (31% of isolates studied) were the most prevalent variant groups (Sharma et al., 2011). Variant group I and II were most common in Portugese, Galicia and Spain (Gouveia et al., 2011; Pesqueira et al.,

2015). In China and Brazil, variant group I was identified as the predominant group (Fajardo et al., 2007; Liu et al., 2013). Genetic diversity study by Turturo et al. (2005) involving GLRaV-3 isolates collected from different varieties from 14 countries sequenced using primers specific to

RdRP, HSP70 and CP regions indicated variant group I as the predominant group. The data obtained in this study showed that group I of GLRaV-3 is predominant in vineyard blocks analyzed in this study (Fig. 22). These results suggest that P.maritimus and/or P.corni could be transmitting variant group I more frequently than other variant groups in Washington vineyards.

Future studies should focus on this aspect for improved understanding of the relation between virus variability and the spread of GLD. Additionally, studies on identifying genes or proteins involved in virus-vector interactions would help to better understand the disease spread and design strategies for minimizing the spread of GLRaV-3. Few reports have shown that RNAi gene silencing could be induced in plants against a specific insect gene, thus controlling the insect populations (Baum et al., 2007; Mao et al., 2007). Injection experiments have shown that the hemipteran insects are sensitive to systemic RNAi (Mutti et al., 2006; Ghanim et al., 2007;

Liu et al., 2010). Induction of RNAi in the grapevine with phloem localized siRNAs that target the gene involved in virus-vector interaction would help to control the vector populations as well as minimize the spread of the virus. Although RNAi involves creating a genetically modified plant that might raise concerns to the wine grape industry, this strategy would prove to be of great value toward overcoming negative impacts of the disease (Skaljaca et al., 2010).

The first line of defense against managing GLD is planting virus tested ‘clean’ cuttings.

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Although virus-tested planting stock obtained from certified nurseries was advocated for planting new vineyards to gain economic benefits (Fuller et al., 2015), our results indicated that apparently healthy vineyards are subjected to the constant risks of disease pressure from neighboring infected vineyards. Therefore, management strategies should be implemented in the field after planting certified grapevine cuttings. In Cabernet Sauvignon block, the results based on the temporal increase of disease and spatial autocorrelation analysis suggested a primary spread of the disease from 2008 to 2011. After 2011, there is a significant spatial autocorrelation due to a dramatic shift in the disease incidence and clustering of infected vines suggesting secondary spread of the disease within the young block. These observations point towards rigorous control measures such as uprooting of infected vines along with vector control implemented prior to the start of the secondary spread for effective control of GLD.

Although, insecticides are widely used to control the vector population of mealybugs, insecticide applications were found to be less effective in reducing vector populations and ensuing viral spread in vineyards (Daane et al., 2012; Almeida et al., 2013; Wallingford et al.,

2015). In this study, although the young vineyard blocks were treated with systemic insecticdes, mealybugs were spotted on a few vines showing leafroll symptoms (data not shown). Thus, treating both old infected and newly planted healthy blocks with appropriate insecticides could help minimizing the movement of vectors from a symptomatic vine to a healthy vine.

Furthermore, GLRaV-3 is shown to persist in the remnant roots from the preceding vineyard

(Bell et al., 2009). Use of systemic insecticides prior to vineyard removal, use of herbicide to prevent the persistence of living Vitis material from a preceding vineyard or allowing a fallow period before re-planting with clean cuttings would reduce the potential of virus transmission by mealybugs. Pheromone-based mating disruption was also shown to be effective in controlling

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mealybug populations at low densities (Walton et al., 2006). Altogether, a combination of vector control together with uprooting of infected vines has proven to be an effective management strategy to control the spread of GLD (Pietersen and Walsh, 2012).

Even though the damage caused by leafroll viruses is not severe enough as to mandate the removal of all infected vines as in other tree diseases (Dallot et al., 2003), the benefits of roguing should not be underestimated in reducing the spread of GLD (Cabaleiro et al., 1999,

2008). Continuous rouging of infected vines in the first few years of disease spread was found to be effective in eradicating the virus from vineyards (Pietersen, 2006). The data obtained from this study will help designing models to analyze the temporal and spatial spread of the disease for optimizing strategies to minimizing the spread of GLD.

In summary, our study on the spatial and temporal dynamics of GLD provided a baseline data to conduct further research for a better understanding of various confounding factors contributing to the spread of GLD across vineyards planted in widely variable geo-climatic conditions of Washington State. Our overall observations of spatio-temporal dynamics of GLD are similar to those reported from other regions (Australia: Habili and Nutter, 1997; South

Africa: Pietersen, 2006; Spain: Cabaleiro et al., 2008; California, USA: Golino et al., 2008; New

Zealand: Charles et al., 2009; Italy: Gribaudo et al., 2009; France: Le Maguet et al., 2013).

Taken together, the general features of spatial and temporal dynamic processes contributing to

GLD epidemics appear to be common across the grapevine-growing regions worldwide.

However, region and site-specific differences with regard to vector species distribution, climatic factors and viticultural practices need to be considered for a comprehensive understanding of

GLD in Washington vineyards relative to other regions.

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ACKNOWLEDGEMENTS

This research was supported, in part, by the WSU Agricultural Research Center, the Wine

Advisory Committee, the Washington Wine Commission and the Washington State Grape & Wine

Research Program. Bhanu Priya Donda is grateful to Altria - Chateau Ste. Michelle Wine Estates for providing graduate research assistantship. We thank Olufemi Alabi, Sridhar Jarugula, Tefera

Mekuria, Andrew Schultz and Eunice Kanuya-Beaver for their help with the data collection during

2007 and 2012.

AFFLIATIONS OF CO-AUTHORS

Dr. Sandya R. Kesoju

United States Department of Agriculture, Irrigated Agriculture Research and Extension Center,

Washington State University, Prosser, Washington, United States of America.

Dr. Neil Mc Roberts

Department of Plant Pathology, University of California, Davis, California, United States of

America.

Dr. Rayapati A. Naidu

Department of Plant Pathology, Washington State University, Irrigated Agriculture Research and

Extension center, Prosser, Washington, United States of America.

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AUTHORS' CONTRIBUTIONS

Conceived and designed the experiments: RAN. Performed the experiments: BP. Analyzed the data: BP, SRK, NMR and RAN. Contributed reagents/materials/analysis tools: RAN. Wrote the paper: BP, SRK and RAN.

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REFERENCES

Alabi, O. J., Gutha, L. R., Casassa, L. F., Harbertson, J., Mirales, M., Beaver, C. W., Davenport, J., and Naidu, R. A. 2012. Impacts of grapevine leafroll disease on own- rooted wine grape cultivar in cool climate conditions. 63rd American Society for Enology and Viticulture National Conference. Portland, OR, U.S.A, pp. 69-70.

Almeida, R. P. P., Daane, K. M., Bell, V. A., Blaisdell, G. K., Cooper, M. L., Herrbach, E., and Pieterson, G. 2013. Ecology and management of grapevine leafroll disease. Front. Microbiol. 4:94. doi:10.3389/fmicb.2013.00094.

Al Rwahnih, M., Dolja, V. V., Daubert, S., Koonin, E. V., and Rowhani, A. 2012. Genomic and biological analysis of grapevine leafroll-associated virus 7 reveals a possible new genus within the family Closteroviridae. Virus Res. 163:302–309. doi:10.1016/j.virusres.2011.10.018.

Atallah, S. S., Gómez, M. I., Fuchs, M. F., and Martinson, T. E. 2012. Economic impact of grapevine leafroll disease on Vitis vinifera cv. Cabernet franc in Finger Lakes vineyards of New York. Am. J. Enol. Vitic. 63:73–79. doi:10.5344/ajev.2011.11055.

Bahder, B. W., Alabi, O. J., Poojari, S., Walsh, D. B., and Naidu, R. A. 2013a. A survey for grapevine viruses in Washington State ‘Concord’ (Vitis x labruscana ) vineyards. Online. Plant Health Progress. doi:10.1094/PHP-2013-0805-01-RS.

Bahder, B. W., Poojari, S., Alabi, O. J., Naidu, R. A., and Walsh, D. B. 2013b. Pseudococcus maritimus (Hemiptera: Pseudococcidae) and Parthenolecanium corni (Hemiptera: Coccidae) are capable of transmitting grapevine leafroll-associated virus 3 between Vitis x labruscana and Vitis vinifera. Environ. Entomol. 42:1292–1298. doi:10.1603/EN13060.

Basso, M. F., Fajardo, T. V. M., Santos, H. P., Guerra, C. C., Ayub, R. A., and Nickel, O. 2010. Leaf physiology and enologic grape quality of virus-infected plants. Trop. Plant Pathol. 35:351– 59. doi: 10.1590/S1982-56762010000600003.

Baum, J. A., Bogaert, T., Clinton, W., Heck, G. R., Feldmann, P., Ilagan, O., Johnson, S., Plaetinck, G., Munyikwa, T., Pleau, M., Vaughn, T., and Roberts J. 2007. Control of coleopteran insect pests through RNA interference. Nat. Biotechnol. 25:1322–6. doi:10.1038/nbt1359.

Bell, V. A., Bonfiglioli, R. G. E., Walker, J. T. S., Lo, P. L., Mackay, J. F., and McGregor, S. E. 2009. Grapevine leafroll-associated virus 3 persistence in Vitis vinifera remnant roots. J. Plant Pathol. 91:527–533.

Bester, R., Maree, H. J., and Burger, J. T. 2012. Complete nucleotide sequence of a new strain of grapevine leafroll-associated virus 3 in South Africa. Arch. Virol. 157:1815–1819. doi:10.1007/s00705-012-1333-8.

Boots, B. N., and Getis. A. 1998. Point pattern analysis. California: Sage Publications.

123

Bovey, R., and Martelli, G. P. 1992. Directory of major virus and virus-like diseases of grapevines. Mediterranean Fruit Crop Improvement Council (MFCIC), Rome pp. 41–51.

Bovey, R., Gartel, W., Martelli, G.P., and Vuitenez, A. 1980. Virus and Virus-like Diseases of Grapevines. Editions Payot, Lausanne.

Cabaleiro, C., Segura, A., and García-Berrios, J. 1999. Effect of grapevine leafroll associated virus 3 on the physiology and must of Vitis vinifera L. cv. Albariño following contamination in the field. Am. J. Enol. Vitic. 50:40–44.

Cabaleiro, C., Couceiro, C., Pereira, S., Cid, M., Barrasa, M., and Segura, A. 2008. Spatial analysis of epidemics of grapevine leafroll associated virus-3. Eur J Plant Pathol. 121:121–130. doi:10.1007/s10658-007-9254-1.

Charles, J. G., Cohen, D., Walker, J. T. S., Forgie, S. A., Bell, V. A., and Breen, K. C. 2006. A review of grapevine leafroll-associated Virus type 3 (GLRaV-3) for the New Zealand Wine industry. Report to New Zealand Wine growers. Hort Research Client Report. 18447:79.

Charles, J. G., Froud, K. J., Brink, R. V. D., and Allan, D. J. 2009. Mealybugs and the spread of grapevine leafroll-associated virus 3 (GLRaV-3) in a New Zealand vineyard. Australas. Plant Pathol. 38:576–583. doi:10.1071/AP09042.

Chooi, K. M., Cohen, D., Pearson, M. N. 2013. Molecular characterisation of two divergent variants of grapevine leafroll-associated virus 3 in New Zealand. Arch Virol. 158: 1597–1602. doi:10.1007/s00705-013-1631-9.

Daane, K. M., Bentley, W. J., and Weber, E. A. 2004. Vine mealybug: A formidable pest spreads throughout California vineyards. Prac. Winery Vineyard Mag. 3:35-40.

Daane, K. M., Almeida, R. P. P., Bell, V. A., Botton, M., Fallahzadeh, M., Mani, M., Miano, J. L., Sforza, R., Walton, V. M., and Zaviezo, T. 2012. Biology and management of mealybugs in vineyards. Pages 271-308 in: Arthropod Management in Vineyards. N. J. Bostanian, R. Isaacs, and C. Vincent, eds. Springer, Dordrecht, the Netherlands.

Dallot, S., Gottwald, T., Labonne, G., and Quiot, J. B. 2003. Spatial pattern analysis of Sharka disease (Plum pox virus strain M) in peach orchards of southern France. Phytopathology 93: 1543–1552.

Engel, E. A., Girardi, C., Escobar, P. F., Arredondo, V., Domínguez, C., Pérez-Acle, T., Valenzuela, P. D. T. 2008. Genome analysis and detection of a Chilean isolate of Grapevine leafroll associated virus-3. Virus Genes. 37: 110–118. doi: 10.1007/s11262-008-0241-1

Engelbrecht, D. J., and Kasdorf, G. G. F. 1985. Association of a closterovirus with grapevines indexing positive for grapevine leafroll disease and evidence for its natural spread in grapevine. Phytopathol. Mediterr. 24:101–105.

124

Fajardo, T. V. M.., Dianese, E. C., Eiras, M., Cerqueira, D. M., Lopes, D. B., Ferreira, M. A. S. V., and Martins, C. R. F. 2007. Variability of the coat protein gene of grapevine leafroll- associated virus 3 in Brazil. Fitopatol. Bras. 32:335–340. doi:10.1590/S0100- 41582007000400008.

Farooq, A. B. U., Ma, Y. X., Wang, Z., Zhuo, N., Wenxing, X., Wang, G. P., and Hong, N. 2013. Genetic diversity analyses reveal novel recombination events in grapevine leafroll-associated virus 3 in China. Virus Res. 171:15-21. doi:10.1016/j.virusres.2012.10.014.

Fuchs, M., Martinson, T. E., Loeb, G. M., and Hoch, H. C. 2009. Survey for the three major leafroll disease-associated viruses in Finger Lakes vineyards in New York. Plant Dis. 93:395– 401. doi:10.1094/PDIS-93-4-0395.

Fuller, K. B., Alston, J. M., and Golino, D. A. 2015. The economic benefits from virus screening: A case study of grapevine leafroll in the North coast of California. Am. J. Enol. Vitic. 66:112-119. doi:10.5344/ajev.2014.14055.

Ghanim, M., Kontsedalov, S., and Czosneck, H. 2007. Tissue-specific gene silencing by RNA interference in the whitefly Bemisia tabaci (Gennadius). Insect Biochem Mol Biol. 37:732-738. doi:10.1016/j.ibmb.2007.04.006.

Goheen, A. C. 1970. In: Frazier, N.W. (Ed.), Grapevine Leafroll. Virus Diseases of Small Fruits and Grapevines. Division of Agricultural Sciences, University of California, pp. 209–212.

Golino, D. A., Sim, S. T., Gill, R., and Rowhani, A. 2002. California mealybugs can spread grapevine leafroll disease. Cal. Agric. 56:196-201.

Golino, D. A., Wolpert, J., Sim, S. T., Benz, J., Anderson, M., and Rowhani, A. 2009a. Virus effects on vine growth and fruit components of three California ‘Heritage’ clones of Cabernet Sauvignon. Pages 243-244 in: Proc. XVI Int. Counc. Study Viruses Virus-Like Dis. Grapevine. Dijon, France.

Golino, D. A., Wolpert, J., Sim, S. T., Benz, J., Anderson, M., and Rowhani, A. 2009b. Virus effects on vine growth and fruit components of Cabernet Sauvignon on six rootstocks. Pages 245-246 in: Extended Abstr. 16th Meet. ICVG. Dijon, France.

Gouveia, P., Santos, M. T., Eiras-Dias, J. E., and Nolasco, G. 2011. Five phylogenetic groups identified in the coat protein gene of grapevine leafroll-associated virus 3 obtained from Portuguese grapevine varieties. Arch. Virol. 156:413–420. doi:10.1007/s00705-010-0878-7.

Goszczynski, D. E., Kasdorf, G. G. F., Pietersen, G., and Van Tonder, H. 1996. Grapevine leafroll-associated virus-2 (GLRaV-2) - mechanical transmission, purification, production and properties of antisera, detection by ELISA. S Afr. J. Enol. Vitic. 17:15-26.

125

Gribaudo, I., Gambino, G., Bertin, S., Bosco, D., Cotroneo, A., and Man- nini, F. 2009. Monitoring the spread of viruses after vineyard replanting with heat-treated clones of Vitis vinifera “Nebbiolo”. J. Plant Pathol. 91:741–744. doi:10.4454/jpp.v91i3.572.

Habili, N., and Nutter, F. W. 1997. Temporal and spatial analysis of grapevine leafroll- associated virus 3 in Pinot Noir grapevines in Australia. Plant Dis. 81:624–628. doi:10.1094/PDIS.1997.81.6.625.

Jooste, A. E. C., Pietersen, G., and Burger, J. T. 2011. Distribution of grapevine leafroll associated virus 3 variants in South African vineyards. Eur. J. Plant Pathol. 131:371–381. doi:10.1007/s10658-011-9814-2.

Komar, V., Vigne, E., Demangeat, G., Lemaire, O., and Fuchs, M. 2010. Comparative performance analysis of virus-infected Vitis vinifera cv. Savagnin rose grafted onto three rootstocks. Am. J. Enol. Vitic. 61:68–73. doi:10.1093/jxb/ers244.

Kulldorff, M. 1997. A spatial scan statistic. Commun. Stat. Theory Meth. 26:1481-1496.

Lee, J., and Martin, R. R. 2009. Influence of grapevine leafroll associated viruses (GLRaV-2 and -3) on the fruit composition of Oregon Vitis vinifera L. cv. Pinot noir: phenolics. Food Chem.112:889–96. doi:10.1016/j.foodchem.2008.06.065.

Lee, J., Keller, K. E., Rennaker, C., and Martin, R. R. 2009. Influence of grapevine leafroll associated viruses (GLRaV-2 and -3) on the fruit composition of Oregon Vitis vinifera L. cv. Pinot noir: free amino acids, sugars, and organic acids. Food Chem. 117:99–105. doi:10.1016/j.foodchem.2008.06.065.

Le Maguet, J., Fuchs, J. J., Beuve, M., Chadoeuf, J., Herrbach, E., and Lemaire, O. 2013. The role of the mealybug Phenacoccus aceris in the epidemic of Grapevine leafroll associated virus- 1 (GLRaV-1) in two French vineyards. Eur. J. Plant Pathol. 135:415–427. doi:10.1007/s10658- 012-0099-x.

Liu, S., Ding, Z., Zhang, C., Yang, B., and Liu, Z. 2010. Gene knockdown by intro-thoracic injection of double-stranded RNA in the brown planthopper, Nilaparvata lugens. Insect Biochem Mol Biol. 40:666-671. doi:10.1016/j.ibmb.2010.06.007.

Liu, M. -H., Li, M. -J., Qi, H. -H., Guo, R., Liu, X. -M., Wang, Q., and Cheng, Y. -Q. 2013. Occurrence of grapevine leafroll-associated viruses in China. Plant Dis. 97:1339-1345. doi:10.1094/PDIS-01-13-0048-RE.

Mahfoudhi, N., Digiaro, M., and Dhouibi, M. H. 2009. Transmission of grapevine leafroll viruses by Planococcus ficus (Hemiptera: Pseudococcidae) and Ceroplastes rusci (Hemiptera: Coccidae). Plant Dis. 93:999–1002. doi:10.1094/PDIS-93-10-0999.

126

Mao, Y. B., Cai, W. J., Wang, J. W., Hong, G. J., Tao, X. Y., Wang, L. J., Huang, Y. P., and Chen, X. Y. 2007. Silencing a cotton bollworm P450 monooxygenase gene by plant mediated RNAi impairs larval tolerance of gossypol. Nat Biotechnol. 25:1307–13. doi:10.1038/nbt1352.

Maree, H. J., Almeida, R. P. P., Bester, R., Chooi, K. M., Cohen, D., Dolja, V. V., Fuchs, M. F., Golino, D. A., Jooste, A. E. C., Martelli, G. P., Naidu, R. A., Rowhani, A., Saldarelli, P., and Burger, J. T. 2013. Grapevine leafroll-associated virus 3. Front. Microbiol. 82:1-21. doi:10.3389/fmicb.2013.00082.

Maree, H. J., Pirie, M. D., Oosthuizen, K., Bester, R., Rees, D. J. G., and Burger, J. T. 2015. Phylogenomic analysis reveals deep divergence and recombination in an economically important grapevine virus. PLoS ONE 10(5): e0126819. doi:10.1371/journal.pone.0126819.

Martelli, G. P., Agranovsky, A. A., Bar-Joseph, M., Boscia, D., Candresse, T., Coutts, R. H., Dolja, V. V., Falk, B. W., Gonsalves, D., Jelkmann, W., Karasev, A. V., Minafra, A., Namba, S., Vetten, H. J., Wisler, G. C., and Yoshikawa, N. 2002. The family Closteroviridae revised. Arch. Virol. 147:2039–2044. doi:10.1007/s007050200048.

Martelli, G. P., Ghanem-sabanadzovic, N. A., Agranovsky, A. A., Rwahnih, M. A, Dolja, V. V., and Dovas, C. I. 2012. Taxonomic revision of the family Closteroviridae with special reference to the grapevine leafroll-associated members of the genus Ampelovirus and the putative species unassigned to the family. J. Plant Pathol. 94:7–19. doi:10.4454/jpp.fa.2012.022.

Martelli, G. P. 2014. Directory of virus and virus-like diseases of the grapevine and their agents. J. Plant Pathol. 96(1S):1–136. doi:10.4454/JPP.V96I1SUP.

Mutti, N. S., Park, Y., Reese, J. C., and Reeck, G. R. 2006. RNAi knockdown of salivary transcript leading to lethality in pea aphid, Acyrthosiphon pisum. J Insect Sci 6:1-7. doi:10.1673/031.006.3801.

Mikona, C., and Jelkmann, W. 2010. Replication of grapevine leafroll-associated virus 7 (GLRaV-7) by Cuscuta species and its transmission to herbaceous plants. Plant Dis. 94:471–476. doi:10.1094/PDIS-94-4-0471.

Naidu, R. A. 2011. Virus Update: The Status of Washington Vineyards. Viticulture and enology extension news-Fall 2011. WSU. http://wine.wsu.edu/research-extension/publications/newsletter/ (Accessed on September 1, 2015).

Naidu, R. A., Rowhani, A., Fuchs, M., Golino, D., and Martelli, G. P. 2014. Grapevine leafroll: a complex viral disease affecting a high-value fruit crop. Plant Dis.98:1172–85. doi:10.1094/PDIS- 08-13-0880-FE.

Naidu, R. A. and Walsh, D. Is ‘grape virus tax’ hitting your pocketbook? Good Fruit Grower. May 15, 2015. Vol. 66, No. 10, pp 10-11.

127

Naidu, R. A., Maree, H. J., and Burger, J. T. 2015. Grapevine leafroll disease and associated viruses: A unique pathosystem. Annu. Rev. Phytopathol. 53:613-34. doi:10.1146/annurev-phyto- 102313-045946.

Nutter, F. W., Jr. and Parker, S. P. 1997. Fitting disease progress curves using EPIMODEL. Pages 24-28 in: Exercises in Plant Disease Epidemiology, L. Francl and D. Neher, eds. APS Press, St. Paul, MN.

Pesqueira, A. M., Cabaleiro, C., and Velasco, L. 2015. Genetic analysis of grapevine leafroll- associated virus 3 population from Galicia, Spain. Plant Pathology. doi:10.1111/ppa.12413.

Pietersen, G. 2006. Spatio-temporal distribution dynamics of grapevine leafroll disease in Western Cape vineyards. Pages 126-127 in: Proc. 15th Int. Counc. Study Viruses Virus-Like Dis. Grapevine. Stellenbosch, South Africa.

Pietersen, G., and Walsh, H. A. 2012. Development of a LAMP technique for the control of grapevine leafroll associated virus type 3 (GLRaV-3) in infected white cultivar vines by Rouging. Pages 50-51 in: Proc. 17th Int. Counc. Study Viruses Virus-Like Dis. Grapevine. Davis, California, USA.

Pietersen, G., Spreeth, N., Oosthuizen, T., Van Rensburg, A., Van Rensburg, M., Lottering, D., Rossouw, N., and Tooth, D. 2013. Control of grapevine leafroll disease spread at a commercial wine estate in South Africa: A case study. Am. J. Enol. Vitic. 64:296-305.

Rayapati, N. A., O’Neil, S., and Walsh, D. 2008. Grapevine Leafroll Disease. WSU Extension Bulletin EB2027E, pp. 20 Available at: http://cru.cahe.wsu.edu/CEPublications/eb2027e/eb2027e.pdf

Ricketts, K. D., Gomez, M. J., Atallah, S. S., Fuchs, M. F., Martinson, T. E., Battany, M. C., Bettiga, L J., Cooper, M. L., Verdegaal, P. S., and Smith, R. J. 2015. Reducing the economic impact of grapevine leafroll disease in California: Identifying optimal disease management strategies. Am. J. Enol. Vitic. 66:138-147. doi:10.5344/ajev.2014.14106.

Sharma, A. M., Wang, J., Duffy, S., Zhang, S., Wong, M. K., Rashed, A., Cooper, M. L., Daane, K. M., and Almeida, R. P. P. 2011. Occurrence of grapevine leafroll-associated virus complex in Napa Valley. PLoS One. 6(10): e26227. doi:10.1371/journal.pone.0026227.

Skaljaca, M., and Ghanim, M. 2010. Tomato yellow leaf curl disease and plant–virus vector interactions. Isr J Plant Sci. 58:103-111. doi:10.1560/IJPS.58.2.103.

Sokolsky, T., Cohen, Y., Zahavi, T., Sapir, G., and Sharon, R. 2013. Potential efficiency of grapevine leafroll disease management strategies using simulation and real spatio-temporal disease infection data. Aust J Grape Wine Res. 19:431-438. doi:10.1111/ajgw.12037.

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Tamura, K., Peterson, D., Peterson, N., Stecher, G., Nei, M., and Kumar, S. 2011. MEGA5: Molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods. Mol. Biol. Evol. 28:2731–2739. doi:10.1093/molbev/msr121.

Thompson, J. D., Higgins, D. G., and Gibson, T. J. 1994. CLUSTAL W: Improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position- specific gap penalties and weight matrix choice. Nucl. Acids Res. 22:4673-4680. doi:10.1093/nar/22.22.4673.

Tsai, C.-W., Chau, J., Fernandez, L., Bosco, D., Daane, K.M. and Almeida, R.P.P. 2008. Transmission of grapevine leafroll-associated virus 3 by the vine mealybug (Planococcus ficus). Phytopath. 98:1093–1098.

Tsai, C. W., Rowhani, A., Golino, D. A., Daane, K. M., and Almeida, R. P. P. 2010. Mealybug transmission of grapevine leafroll viruses: an analysis of virus–vector specificity. Phytopathology 100:830–834.

Turturo, C., Saldarelli, P., Yafeng, D., Digiaro, M., Minafra, A., Savino, V., and Martelli, G. P. 2005. Genetic variability and population structure of grapevine leafroll-associated virus 3 isolates. J. Gen. Virol. 86:217–224. doi:10.1099/vir.0.80395-0.

Wallingford A.K., Fuchs M.F., Martinson T., Hesler S. and Loeb G.M., 2015. Slowing the spread of grapevine leafroll associated viruses in commercial vineyards with insecticide control of the vector, Pseudococcus maritimus (Erhorn) (Hemiptera: Pseudococcidae). J. Insect Sci. 15:112. doi:10.1093/jisesa/iev094.

Wang, J., Sharma, A. M., Duffy, S., and Almeida, R. P. P. 2011.Genetic diversity in the 3’ terminal 4.6-kb region of Grapevine leafroll associated virus 3. Phytopathology 101:445–450.

Walton, V. M., Daane, K. M., Bentley, W. J., Millar, J. G., Larsen, T. E., and Malakar-Kuenen, R. 2006. Pheromone-based mating disruption of Planococcus ficus (Hemiptera: Pseudococcidae) in California vineyards. J. Econ. Entomol. 99:1280–1290. doi:10.1093/jee/99.4.1280.

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CHAPTER FOUR

SUMMARY AND CONCLUSIONS

Grapevine leafroll disease (GLD) is recognized as one of the most serious impediments to long-term sustainability of the grape and wine industry in Washington State. Several virus species, collectively designated as Grapevine leafroll-associated viruses (GLRaVs; family:

Closteroviridae), have been found associated with GLD worldwide. Among them, GLRaV-1, -2,

-3, and -4 and its strains GLRaV-5 and GLRaV-9 were documented in Washington vineyards.

Due to its economic importance, management of GLD is a high priority for Washington’s grape and wine grape industry. A comprehensive understanding of the disease and associated viruses will help deploying science-based disease management strategies for sustainable growth of the grape and wine grape industry in Washington State. Towards this objective, studies were conducted to better understand the molecular biology and epidemiology of viruses associated with GLD.

In the first study, the complete genome sequence of Grapevine leafroll-associated virus 1

(GLRaV-1), the second most widely distributed virus across many grapevine-growing regions, was determined. The genome of GLRaV-1 isolates WA-CH and WA-PN was determined to be

18,731 (WA-CH; Accession number: KU674796) and 18,946 (WA-PN; Accession number:

KU674797) nucleotides, respectively. Based on the sequence data, it appears that the genome of

GLRaV-1 is the second largest, after Citrus trizteza virus, among members of the family

Closteroviridae. The overall genome organization of GLRaV-1 isolates WA-CH and WA-PN with hallmark gene array consisting of nine open reading frames (ORFs) is similar to GLRaV-1 isolates reported previously from Australia and Canada. However, ORF-by-ORF comparisons

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indicated that GLRaV-1 isolates from Washington are more closely related to each other than either of them to the Canadian and Australian isolates. Besides, distinct differences in the size and nucleotide sequence composition were observed in the 5’ and 3’ non-translated region

(NTR) of the two Washington isolates. In Northern blots, three of the eight 3’co-terminal subgenomic RNAs (sgRNAs) were detected at higher levels and were putatively designated as sgRNAs specific to CP, p21 and p24. Among them, the sgRNA corresponding to p24 gene accumulated at the highest level, followed by sgRNAs for CP and p21, respectively. The 5’ termini of five putative sgRNAs were mapped and their leader sequences determined. Using the

5’NTR sequences of WA-CH and WA-PN isolates, reverse-transcription-PCR based restriction fragment length polymorphism assay was developed to distinguish GLRaV-1 variants in vineyards. The results from this study will provide a foundation for the development of an amenable reverse genetic system in the future for elucidating the comparative molecular biology of grapevine-infecting members of the family Closteroviridae.

Although studies on the epidemiology of GLD has been conducted in different grapevine- growing regions in the US, South Africa, Australia, New Zealand and many European countries, very little research has been carred out on field spread of GLD in Washington vineyards.

Previous studies have indicated that GLRaV-3 is more widespread across Washington vineyards and is predominantly associated with GLD. Growers have been advised to use virus-tested, certified planting stock for planting new vineyard blocks as a first line of defense to minimize the spread of GLD. However, the risk of GLD spread into new plantings remains a significant concern for wine grape growers in Washington State. In this study, the spread of GLD was monitored in vineyard blocks planted with certified, virus-tested cuttings of three wine grape cultivars adjacent to heavily infected old blocks. During each season, the position of vines

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showing GLD symptoms was recorded in a matrix representing the planting lattice. Viral status of symptomatic vines was verified by molecular diagnostics assays and found to be positive only for Grapevine leafroll-associated virus 3 (GLRaV-3, genus Ampelovirus), the most ubiquitous virus associated with GLD in Washington vineyards. Temporal analysis of the disease incidence indicated differences in the rate of spread among the three cultivars. Spatial mapping of the disease in young blocks over successive seasons showed a disease gradient in which the highest percentage of symptomatic vines were in rows proximal to heavily infected old blocks. Spatial autocorrelation suggested random distribution of symptomatic vines during initial years indicating primary spread and clustering of symptomatic vines during subsequent years suggestive of secondary spread within a block. Sequence analysis of a portion of the heat-shock protein 70 homolog gene encoded by GLRaV-3 revealed the predominance of one of the genetic variants of the virus in young vineyard blocks. The results provided for the first time science- based knowledge on the spread of GLD in young vineyards to implement site-specific disease management strategies for mitigating negative impacts of the disease under conditions prevailing in Washington State.

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