GENETIC BASIS for HOST RESPONSE to HOP STUNT VIROID by JEFF MARTIN BULLOCK a Dissertation Submitted in Partial Fulfillment of Th

Home , Hop stunt viroid, Hostuviroid, Pospiviroidae, Viroid

GENETIC BASIS FOR HOST RESPONSE TO HOP STUNT VIROID

JEFF MARTIN BULLOCK

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

DOCTOR OF PHILOSOPHY

WASHINGTON STATE UNIVERSITY Department of Plant Pathology

MAY 2016

To the Faculty of Washington State University:

The members of the Committee appointed to examine the dissertation of JEFF MARTIN BULLOCK find it satisfactory and recommend that it be accepted.

______Kenneth C. Eastwell, Ph.D., Chair

______Hanu R. Pappu, Ph.D.

______Brenda K. Schroeder, Ph.D.

______Paul D. Matthews, Ph.D.

ACKNOWLEDGEMENTS

I have been very fortunate to have Dr. Kenneth C. Eastwell as my advisor and mentor throughout this process. His guidance has been invaluable and his efforts on my behalf have been extraordinary! I was one of his first graduate students working on a master’s degree in

1986, now 30 years later I will be his last graduate student to complete a Ph.D. under his guidance. I cannot think of a better person to have guided my path, he has been a huge influence on my scientific training.

I would also like to thank my committee members, Dr. Hanu R. Pappu, Dr. Brenda K.

Schroeder, and Dr. Paul D. Matthews for their constant encouragements, technical assistance and support.

In addition I am grateful to the Washington Hop Commission and the Hop Research

Council for funding this project and to all the members of the Northwest Clean Plant Center, namely: Jan Burgess, Shannon Santoy, Tina Vasile, Syamkumar Sivasankara, Piotr Kowalec,

Debbie Woodbury, Dan Villamor, Holly Ferguson, and Eunice Beaver-Kanuya for all their help and reassurances. I would like to especially thank Martin Joseph for his assistance in plant propagation from tissue culture to greenhouse maintenance and Madhu Kappagantu, my fellow graduate student, for her help with plant care and her important insights in bioinformatics.

Most importantly I would like to thank my parents Gil and Jean Bullock, and my wife Jan for their ever present love and unwavering support, without them this goal could not have been reached!

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GENETIC BASIS FOR HOST RESPONSE TO HOP STUNT VIROID

Abstract

by Jeff Martin Bullock, Ph.D. Washington State University May 2016

Chair: Kenneth C. Eastwell, Ph.D.

Hop stunt viroid (HSVd) was first detected in hop orchards in Washington State in 2004.

A 2012 survey for the presence of HSVd, in three major hop growing regions in Washington

State indicated an overall incidence of 17.3 percent. A five year yield study comparing HSVd infected and uninfected hop cultivars ‘Glacier’ and ‘Nugget’ revealed that, of those tested,

‘Nugget’ was the most tolerant hop cultivar to HSVd infection and ‘Glacier’ was the most severely affected.

In this study, whole genome and RNA sequence analysis identified potential genes associated with expression of hop stunt disease, and identified DNA-markers that distinguish

HSVd-tolerant plants from HSVd-sensitive cultivars. Healthy plants and HSVd-infected ‘Glacier’ and ‘Nugget’ revealed different gene expression profiles that suggested that a pathogenesis- related protein, thaumatin-like protein (TLP), is a candidate marker for HSVd sensitivity.

TLP was observed to be down regulated 2.6 fold in ‘Glacier’ infected with HSVd compared to HSVd-free ‘Glacier’; no change in TLP expression was observed in ‘Nugget’

iv between HSVd-infected or HSVd free plants. RNA sequence analyses were confirmed using qRT-PCR for TLP expression levels normalized to Glyceraldehyde 3-phosphate dehydrogenase

(GAPDH).

Alignments of the TLP coding sequence (CDS) and the region immediately up-stream of the TLP CDS for each cultivar revealed no sequence variations 100 bp upstream, which includes the ATG translation start site, TATA box and the transcriptional start site. However, 23 nucleotide variations exist between ‘Glacier’ and ‘Nugget’ in a 397 bp region up stream of the first CAAT cis-regulatory element (CRE). These variations create eleven unique CRE differences between ‘Glacier’ and ‘Nugget’. Additionally, within the TLP CDS, nineteen single nucleotide polymorphisms (SNPs) were observed in ‘Glacier’ relative to ‘Nugget’. Several of these SNPs result in amino acid substitutions. Four of the SNPs within the ‘Glacier’ TLP CDS result in amino acid substitutions that were not observed in five other cultivars of hop. In total, 42 SNPs were identified that potentially may be used in marker assisted breeding programs to distinguish potential HSVd-tolerant plants from HSVd-sensitive plants. Evaluation of additional hop genotypes will be necessary to confirm this relationship.

TABLE OF CONTENTS

Page

ACKNOWLEDGEMENTS iii

ABSTRACT iv

LIST OF FIGURES viii

LIST OF TABLES xi

LIST OF ABBREVIATIONS xiii

CHAPTER

1. GENERAL INTRODUCTION 1

Taxonomy and breeding history of genus Humulus 1

Hop compounds essential to brewing 6

Viroids 17

Operons 28

Gene regulation 30

Small RNAs 36

Pathogenesis related proteins 43

Summary and Project goals 45

2. MATERIALS AND METHODS 49

Cloning 50

Cucumber and hop inoculations 53

RNA extractions and high through-put sequencing 59

RNA sequence expression analysis and gene discovery 61

Sequence variations between hop cultivars 66

Relative HSVd concentration and TLP expression levels 68

Survey for the presence of HSVd 69

RNA extraction from FTA cards and RT-PCR for the presence of HSVd 70

3. RESULTS and DISCUSSION 71

Infectious clone of HSVd 71

High through-put sequencing 79

TLP discovery and expression differences between treatment groups 85

Relative HSVd concentration and TLP expression levels 89

TLP coding sequence and sequence analysis 91

Plant specific TLP isoforms 93

Sequence variations in the TLP CDS and regions upstream 96

Survey for the presence of HSVd in Washington State 103

Discussion 104

References 119

vii

LIST OF FIGURES

CHAPTER 1

1 Leaf pattern common to Humulus lupulus and Humulus japonicas. 2

2 Female seed cones (strobiles), female inflorescences, and male inflorescences. 4

3 Female hop cone showing bracts, bracteoles, and cutaway view of hop cone exposing lupulin glands. 8

4 Pathway for humulone synthesis using the branched chain acyl-CoA thioester; isovaleryl-CoA, as a precursor in the presence of valerophenone synthase (VPS) to produce the first intermediate, phlorisovalerophenone (PIVP), in the alpha acid pathway. 10

5 The acyl-CoAs, isovaleryl-CoA, isobutyryl-CoA and 2-methylbutyryl-CoA used in alpha and beta acid synthesis are break down products produced by the catabolism of the branched-chain amino acids (BCAA) leucine, valine and isoleucine. 11

6 BCAA biosynthesis from pyruvate in plants. Valine and isoleucine are produced via parallel pathways; both use identical enzymes. 14

7 Multibranched secondary structure showing hairpins, pseudoknots, and loops that form hammerhead ribozymes, which are unique to Avsunviroidae viroids; less complex secondary rod like structure of Pospiviroidae viroids. 21

viii

CHAPTER 3

1 Capillary gel electrophoresis of PCR amplification products using plasmid F56 purified from transformed One Shot Cells as a template. 71

2 Sequence of F56 plasmid. 72

3 Sequence of monomer from F56 plasmid aligned to hKFKi a well characterized HSVd isolated from hop. 73

4 Mock inoculated cucumber (Cucumis sativus L. cv. Suyo Long). HSVd (F56) inoculated cucumber (Cucumis sativus L. cv. Suyo Long). 74

5 Capillary gel electrophoresis of RT-PCR amplification products using total RNA extracted from cucumber leaf tissue. 75

6 Sequence alignment of HSVd RT-PCR product produced using total RNA extracted from F56 inoculated cucumber as a template and HSVd isolate hKFKi.76

7 ‘Glacier’ plants 140 days post inoculation. 77

8 ‘Nugget’ plants 140 days post inoculation. 78

9 ‘Nugget’ and ‘Glacier’ plants 140 days post inoculation. 79

10 Leaves mock inoculated and HSVd inoculated. 80

11 RNA extraction resolved on a 1% non-denaturing agarose gel. 81

12 Alignment of 58 bp region of contig LA331881 to Brassica napus thaumatin-like protein and alignment of LA331881 putative TLP region to scaffold LD139544. 85

13 Simple linear regression analysis of qRT-PCR results showing the correlation between normalized TLP levels and HSVd levels relative to an HSVd standard for ‘Glacier’ and ‘Nugget’ HSVd infected plants. 90

14 Identified coding sequence from LD139544 region 29612 to 30283 91

15 Translation product, 223 amino acids from identified open reading frame using LD139544, region 29612 to 30283 and PBLAST analysis identified the amino acid sequence as a TLP specific to the plant subfamily. 92

16 Sequence 200 bp upstream of the TLP ATG translation start site 93

17 Amino acid sequence of different plant specific TLP isoforms 94

18 Nineteen single nucleotide polymorphisms in ‘Glacier’ relative to the hop cultivar ‘Nugget’ 97

19 Thaumatin amino acid variations between hop cultivars ‘Nugget’, ‘Glacier’ ‘Cascade’, ‘Columbus’, ‘Galena’, and ‘Willamette’ 98

20 Positions of single nucleotide substitutions and multi nucleotide insertions and position of TATA box and CA transcriptional start within the 41 bp GC rich region and cis-regulatory elements unique to ‘Glacier’ and ‘Nugget’ 100

LIST OF TABLES

CHAPTER 1

1 Representative sample of viroids, their known host and the associated viroid family Pospiviroidae or Avsunviroidae 19

CHAPTER 2

1 Hop rooting media. 58

CHAPTER 3

1 Relative fold change between BLAST identified sequences in HSVd infected and mock inoculated plants from ‘Nugget’ and ‘Glacier’ treatment groups. 82

2 Relative fold change for TLP between, HSVd infected and mock inoculated (HSVd free) plants from ‘Glacier’ and ‘Nugget’ treatment groups. 86

3 Results from qRT-PCR, normalized to GAPDH showed no change between HSVd infected and mock inoculated ‘Nugget’ plants. 87

4 Results from qRT-PCR, normalized to GAPDH showed a 2.6 fold change (down regulation) for HSVd infected ‘Glacier’ plants compared to mock inoculated plants. 88

5 Concentration levels of HSVd in infected tissue correlated to TLP expression levels, normalized to GAPDH using qRT-PCR. 89

6 List of scaffolds with regions containing sequence similarity to TLP CDS. 93

7 Relative fold change for different TLP isoforms between, HSVd infected and HSVd free plants from ‘Glacier’ and ‘Nugget’ treatment groups using RPKM values 95

8 Amino acid substitutions in ‘Glacier’ thaumatin primary protein sequence relative to ‘Nugget’ including amino acids at these positions for the hop cultivars ‘Galena’, ‘Cascade’, ‘Columbus’, and ‘Willamette’ 99

9 List of cis-regulatory elements common to ‘Glacier’ and ‘Nugget’ 102

10 Survey results for the presence of HSVd separated by cultivar from the three major hop growing regions in Washington State. 103

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LIST OF ABBREVIATIONS

(ABA) Abscissic acid (ADFVd) Apple dimple fruit viroid (AFCVd) Apple fruit crinkle viroid (AG) AGAMOUS (AGO) Protein ARGONAUTE (AP-2) Activator protein 2 (ASBVd) Avocado sun blotch viroid (ASSVd) Apple scar skin viroid (atp5cdh) Arabidopsis thaliana Delta-1-pyrroline-5-carboxylate dehydrogenase (avr) Avirulence protein (BCAA) Branched-chain amino acids (BCAT) Branched-chain aminotransferase (BLAST) Basic Local Alignment Search Tool (bp) Base pair (BR) Brassinosteroid (BREd) Downstream TFIIB recognition element (BREu) Upstream TFIIB recognition element (C) Centigrade (CBCVd) Citrus bark cracking viroid (CBLVd) Citrus bent leaf viroid (CCCVd) Coconut cadang-cadang viroid (CCHMVd) Chrysanthemum chlorotic mottle viroid (CCR) Central conserved region

xiii

(cDNA) Complementary DNA (CDS) Coding sequence (CEVd) Citrus exocortis viroid (CHS) Chalcone synthase (CK) Cytokinin (CoA) Coenzyme A (COI1) Coronatine insensitive 1 (CP) Core promoter (CPCNW) Clean Plant Center Northwest (CPE) Core promoter elements (CRE) Cis-regulatory elements (CSVd) Chrysanthemum stunt viroid (CTAB) Cetyltrimethylammonium bromide (DCL) Dicer-like protein

(ddh2o) Double distilled water (DIBOA) 2,4-dihydroxy-1,4-benzoxazin-3-one (DNA) Deoxyribonucleic acid (DPE) Downstream promoter element (dsRNA) Double stranded RNA (E. Coli) Escherichia coli (EDTA) Ethylenediaminetetraacetic acid (EIN) Ethylene insensitive (ERF)Eukaryotic release factor (EST) Expressed sequence tags (ET) Ethylene (GA) Gibberellin

xiv

(GAPDH) Glyceraldehyde-3-Phosphate Dehydrogenase (GTFS) General transcription factors (GTP) Guanosine-5'-triphosphate (HBV) Hepatitis B virus (HDAG) Hepatitis D antigen (HDV) Hepatitis delta virus (hlbcat1) Humulus lupulus BCAT1 (hlbcat2) Humulus lupulus BCAT2 (HLVd) Hop latent viroid (hpRNA) Hairpin RNA (HR) Hypersensitive response (HSVd) Hop stunt viroid (Inr) Initiator (IRES) Internal ribosome entry sites (IR-PTGS) IR post-transcriptional gene silencing (JA) Jasmonic (JAR1) Jasmonate resistant 1 (JIN1) Jasmonate insensitive 1 (LB) Luria broth (leexp2) Lycopersicon esculentum Expansin2 (MEP) Methyl-D-erythritol 4-phosphate (miRNA) MicroRNA (ml) Milliliter (mm) Millimeter (Mm) Millimolar (MOPS) 3-(N-morpholino) propanesulfonic acid (Mot1) Modifier of transcription 1

(mRNA) Messenger RNA (MTE) Motif ten element (nat-siRNA-ATGB2) Natural antisense transcript siRNAs ATGB2 (nat-siRNAs) Natural antisense transcript siRNAs (NC2) Negative cofactor 2 (NCBI) National Center for Biotechnology Information (NDR1) Nuclear Dbf2-related kinase 1 (ng) Nanogram (nt) Nucleotide (P) Pathogenicity

(P23) Pathogenesis-related protein 23 (PCR) Polymerase chain reaction (PCR) Polymerase chain reaction (PIC) Pre-initiation complex (PIVP) Phlorisovalerophenone (PLMVd) Peach latent mosaic viroid (pol II) RNA polymerase II (PP2) Phloem protein 2

(PPRL) Pentatricopeptide repeat-like protein gene

(PR) Pathogenesis-related protein (Pst-avrrpt2) Pseudomonas syringae avr protein Rpt2 (PSTVd) Potato spindle tuber viroid (PTGS) Posttranscriptional gene silencing (PVP 40) Polyvinylpyrolidone M. W. 40,000 (qRT-PCR) Quantitative reverse transcription-polymerase chain reaction (R) Resistance protein (Rab2-like) Ras-related protein

xvi

(RDDM) RNA-directed DNA methylation (RDR) RNA dependent RNA polymerase (RNA) Ribonucleic acid (RNAP) RNA polymerase (RNAseq) RNA sequencing (RPKM) Reads Per Kilobase of transcript per Million mapped reads (RPM) Revolutions per minute (RPS2) Resistance to Pseudomonas syringae protein 2 (Rpt2) Root phototropism protein 2 (RT-PCR) Reverse transcription-polymerase chain reaction (RT-qPCR) Reverse transcription quantitative real-time PCR (SAR) Systemic acquired resistance (SDS) Sodium dodecyl sulfate (SEP3) Developmental protein SEPALLATA 3 (siRNAs) Small interfering RNAs (SMZ) Protein SCHLAFMUTZE (SNP) Single nucleotide polymorphisms (SNZ) Protein SCHNARCHZAPFEN (SOC) Super Optimal broth with Catabolite repression (SOC1) SUPPRESSOR OF OVEREXPRESSION OF CONSTANS1 (SRO5) Polymerase SIMILAR TO RCD ONE 5 (TAE) Tris-base, acetic acid and EDTA (Taq) Thermus aquaticus (TASVd) Tomato apical stunt viroid (TBP) TATA box binding protein (TCDVd) Tomato chlorotic dwarf viroid (TCH) Terminal conserved hairpin

xvii

(TCR) Terminal conserved region

(TF) Trans-acting transcription factors (TFII) Transcription factor for RNA polymerase II (TGS) Transcriptional gene silencing (TL) Terminal left

(TLP) Thaumatin like protein (TMV) Tobacco mosaic virus (TOE 1, 2, 3) Protein target of EAT 1, 2, 3

(TR) Terminal right

(Tris) Tris (hydroxymethyl) aminomethane (TSS) Transcriptional start site (UTR) Untranslated region (UV) Ultraviolet (V) Variable

(VPS) Valerophenone synthase (WTSS) Whole transcriptome shotgun sequencing (XCPE1) X core promoter element 1

(l) Microliter

xviii

This dissertation is dedicated to the Spark in all of us!

Casually Fishing the Day

The day rolls around us Like a river we stand in, Knee deep, casting our Bait into its passing.

We expect its coming to continue, excusing itself politely past us, on its way to tomorrow and next year.

We attend to what might be Living within it, fishing for Some magic we can’t anticipate, but are we extracting the minutes

From the hours, the hours from the morning, the morning from the day, Its ordinariness; the guarantee we believe that arrives with it,

Blurs its passage, like an analgesic eases the pang and catch. Are we paying attention? Are we embracing the day?

Spark Dorsey

xix

CHAPTER 1

INTRODUCTION

Taxonomy and breeding history of genus Humulus.

Humulus lupulus var. lupulus (common hop or hop) used in the brewing industry is a flowering perennial dioecious herb which grows new shoots each spring from over wintering rhizomes and dies away in late fall leaving a cold tolerant rhizome below the soil surface. Botanically, hops are classified as bines as opposed to vines. The difference being bines support themselves by shoot growth and vines use tendrils or suckers to climb. Taxonomically, Humulus is in the family Cannabaceae, which contains only ten genera including; Humulus and Cannabis (Yang et al., 2013). Three species have been identified in the Humulus genus: H. lupulus L., H. japonicus

Siebold and Zucc, also known as H. scandens (Lour.) Merr. and H. yunnanensis (Small, 1978). H. lupulus and H. yunnanensis have a leaf pattern that contains 3-5 lobes, while that of H. japonicas has 5-7 lobes (Figures 1 A and B). Features that differentiate H. lupulus and H. yunnanensis include: leaf surface, pollen size, gland size, and number and dissemination of trichomes (Small, 1978; Small 1980). Additionally, H. yunnanensis is concentrated within southern China within the Yunnan province and is rarely found in other hop growing regions throughout the world.

H. lupulus is agriculturally the most important of the three species. Taxonomic varieties within

H. lupulus include: H. lupulus var. lupuloides E. Small (syn. H. americanus Nutt.), H. lupulus var. lupulus, H. lupulus var. cordifolius (Miquel) Maximowicz, H. lupulus var. pubescens E. Small, and

H. lupulus var. neomexicanus Nelson and Cockerell. Many of the cultivars used in the current brewing industry are offspring from hybrid crosses of H. lupulus var. lupulus, a European hop, and one of the north American hop varieties H. lupulus var. lupuloides or H. lupulus var. neomexicanus.

A B

Figure 1. A) Three to five lobe leaf pattern common to Humulus lupulus. B) Five to seven lobe leaf pattern common Humulus japonicas. (Photos courtesy of Wikimedia Commons,

the free media repository https://commons.wikimedia.org/wiki/wikimedia.org)

Historical evidence suggests that Humulus lupulus, was first cultivated several thousand years before the Christian era in central and western Asia (Unger, 2004). Before the 8th century AD the primary use of hops was for medicinal purposes not brewing (Edwardson, 1952). It is uncertain when hops were first used in the brewing process, but the first written reference for the use of hops as a flavoring agent in fermented alcoholic drinks dates to AD 822 (Unger,

2004). The first organized breeding program designed to improve hop varieties for the purpose of brewing started at the turn of the 20th century in 1904 (Neve, 1986).

Like the domestication of other agriculturally important crops, early breeding of hops was via open pollination and selection of new plants with specific desirable traits. Only female plants produce seed cones, also known as strobiles, that are used in the brewing process (Figure 2A).

However, both male and female plants produce inflorescences (Figures 2B-C). Early cultivars of hops were often named for the geographical area from which they were cultivated, e.g.,

‘Tettnanger’, named for the Tettnang area of Germany. Other early cultivars produced via open pollination and grower selection include: ‘Golding’ and ‘Fuggle’ from England, and ‘Styrian

Golding’ and ‘Saaz’ from Europe. These early varieties were all the result of hop growers selecting and cultivating plants from their hop gardens rather than an organized program to produce new hop varieties. Many of these, such as ‘Golding’ were “improved clones” rather than plants from a seedling such as ‘Fuggle’ (Neve, 1986). Only female plants are actively cultivated in North America and male plants are used for breeding purposes only.

B C

Figure 2. A) Female seed cones (strobiles). B) Female inflorescences. C) Male inflorescences. (Photos courtesy of Wikimedia Commons, the free media repository https://commons.wikimedia.org/wiki/wikimedia.org)

The first organized breeding program designed to improve hop varieties for the purpose of brewing started at the turn of the 20th century in 1904 at Wye College in Kent, England under the guidance of E.F. Salmon (Neve, 1986). Salmons' objectives were to create new varieties of hop plants that were resistant to the most prevalent pest and diseases, yielded cones with high concentrations of alpha and beta acids, and had good aroma (Neve, 1986; Patzak, 2005).

Because of their high concentrations of alpha and beta acids, Salmon introduced wild North

American hops into his breeding program (Neve, 1986). The first two successful varieties from

Wye College were, ‘Brewer's Gold’ and ‘Bullion’ released in 1934 and 1938, respectively. Both had high concentrations of alpha and beta acids, but undesirable aroma. A later hybrid released in 1944, ‘Northern Brewer’ produced a good aroma and also had high yields of alpha and beta acids (Neve, 1986).

Early cultivars of European hops are diploid having nine pairs of autosomal chromosomes in male and female plants and two X chromosomes in female and an X, Y combination in males

(Seefelder et. al., 2000). However tetraploid plants are occasionally found in natural populations and tetraploid plants are easily produced by treating diploids with colchicine

(Haunold, 1972; Beatson, 1993). In many current breeding programs, tetraploid males are crossed with diploid females to produce triploids. Triploids are often sterile, produce few seeds and are associated with higher alpha and beta acid yields and increased plant vigor

(Beatson, 2003).

In an early study (Haunold, 1971) a tetraploid derived from a colchicine-treated ‘Fuggle’ female was crossed with six different, but related ‘Fuggle’ males, each of which were diploid.

Results from this cross showed seedlings were 76.3 % triploids, 22.4% aneuploids, and the remainder tetraploids or diploids. The triploids had greater vigor and higher cone yields compared to the diploid parents or non-triploid offspring. For example, two year-old triploids over the same growing period had an average growth of 121 mm/day compared to 53 mm/day and 103mm/day for tetraploid and diploid plants, respectively (Haunold, 1971).

Additionally the triploids average cone weight per plant was 1.7 to 4.6 times greater in mass with less seed weight than any of the diploid and tetraploid parents or offspring from the same cross. This additional cone weight was associated with a greater number of resin or lupulin glands producing higher amounts of the bittering agents and essential oils that brewers are most concerned with (Haunold, 1971).

Hop compounds essential to brewing.

Brewer masters are predominantly interested in secondary metabolites produced in the resin or lupulin glands (Figure 3) found in high concentration on the cones of female plants. The glands are also present on male and female leaves and other tissues, but not in great quantity

(De Keukeleire et al., 2003). The characteristic bitter taste in beer is produced by alpha and beta acid present in the resins, and the hoppy and other aroma flavors are dependent on essential oils. Both the resins and essential oils are themselves complex mixtures of compounds.

European varieties are better known for their aroma qualities, while North American varieties are generally known for their bittering agents. As a result of successful breeding programs, a

6 great number of hops grown for commercial brewing are hybrids of European and North

American varieties. Regardless of the varieties used to create a new cultivar, hops are frequently placed into one of two categories with multiple names for each: kettle, bitter, or high-alpha hops are used for beer bittering, and aroma, noble, or European hops are used more for flavoring (Verzele and De Keukeleire, 1991).

Cones of all hops consist of overlapping bracts and bracteoles (Figure 3A). The dry weight of cones is mostly cellulose, approximately 40%, while lupulin gland mass accounts for 20%, and the remainder is proteins, tannins, waxes, water, pectins, and other complex carbohydrates

(Howard, 1964). According to various chemical and physical properties, hop secondary metabolites from lupulin glands (Figure 3B) can be categorized into three distinct types: resins, oils, and polyphenols. Hop hard and soft resins are soluble in cold methanol and diethyl ether.

Differences in solubility of hard and soft resins exist in hexane: soft resins are soluble, hard resins are not. Soft resins contain the alpha and beta acids, and hard resins contain flavanones and prenylated chalcones, with xanthohumol being the most abundant. Essential oils are traditionally compounds that can be separated by steam distillation. In hop, these largely include: alpha-humulene, monoterpene, myrcene, beta-caryophyllene, and other organic compounds such as the sesquiterpenes. Also found in lupulin of hop cones are flavonol glycosides, aromatic carboxylic acids, and non-prenylated flavonoids (Steenackers, et al., 2015).

An understanding of the brewing properties of both the beer bittering and flavoring compounds which constitute lupulin has provided breeders with useful chemical measurements for assessing promising new cultivars. These initially included the major components,

7 humulone, cohumulone, and adhumulone, which comprise up to 95% of the alpha acid fraction and lupulone, colupulone, adlupulone, which compose the beta acids (Stevens, 1967). Often a hops quality or potential will be assessed by the weight percent of alpha acids in the cones.

Such a high importance is placed on alpha acids that hop alpha acid production is used in the world hop market analysis by the world's largest supplier of hops and hop products

(Georgensgmuend, 2013). It is well recognized that alpha acid content and the ratio of each type are varietal characteristics (Meilgaard, 1960; Nickerson et al., 1986).

B A

Figure 3. A) Female hop cone showing bracts and bracteoles. B) Cutaway view of hop cone exposing lupulin glands, magnified 40 X. (Photos courtesy of Wikimedia Commons, the free media repository https://commons.wikimedia.org/wiki/wikimedia.org)

While there is a high level of agreement on beer bittering compounds, there is less agreement on the basis of beer aromas. Hundreds of chemical substances have been recognized as hop essential oils, but only a few have been established in beer. Oil configuration varies with variety, brewing procedures and recipes. Components can evaporate, oxidize, and undergo isomerization and other types of chemical transformations at any stage during the brewing or harvesting process (Neve, 1991). Consequently, detection and quantification of essential oils in green hop cones is less likely to be used as good measurements for assessing promising new cultivars.

The biochemistry and enzymatic pathways relevant to alpha acid production are well understood. Alpha and beta acids produced by hops are prenylated acylphloroglucinols. The primary alpha acids include humulone, adhumulone, and cohumulone (Verzele and De

Keukeleire, 1991) but several more have been identified (Briggs, et al., 2000). Iso-alpha acids are isomeric derivatives of the alpha acids that are produced as the result of alpha-acid isomerization during the brewing process and are the primary bittering acids in beer (Verzele,

1986). Zuurbier et al. (1995) proposed the first complete biosynthetic pathway for humulone and cohumulone using the branched chain acyl-CoA thioesters: isovaleryl-CoA, isobutyryl-CoA, and malonyl-CoA as precursors in the presence of chalcone synthase (CHS) to produce the first intermediate, phlorisovalerophenone (PIVP), in the humulone and cohumulone pathway

(Figure 4).

Figure 4. Pathway for humulone synthesis using the branched chain acyl-CoA thioester; isovaleryl-CoA, as a precursor in the presence of valerophenone synthase (VPS) to produce the first intermediate, phlorisovalerophenone (PIVP), in the alpha acid pathway. Catabolism

of Leucine which provides the initial building blocks for the production of isovaleryl-CoA is physically separated in cells from branched chain amino acid (BCAA) synthesis. BCAA catabolism in plants is localized to mitochondria. (Adapted from Clark et al., 2013.)

It was shown latter that a member of the CHS superfamily, valerophenone synthase (VPS) was used in hop to produce PIVP (Paniego et al., 1999). The primary amino acid sequence for VPS expressed in hop has been determined and consists of 394 amino acids (Okada and Ito, 2001).

The acyl-CoAs, isovaleryl-CoA, isobutyryl-CoA and 2-methylbutyryl-CoA (Figure 5), used in alpha and beta acid synthesis are break down products produced by the catabolism of the branched- chain amino acids (BCAA) leucine, valine and isoleucine, respectively (Singh and Shaner, 1995).

Figure 5. The acyl-CoAs, isovaleryl-CoA, isobutyryl-CoA and 2-methylbutyryl-CoA used in alpha and beta acid synthesis are break down products produced by the catabolism of the branched-chain amino acids (BCAA) leucine, valine and isoleucine. (Adapted from Clark et al., 2013.)

These amino acids are considered essential amino acids as they are not produced by humans.

In plants it is believed they are produced in the chloroplast (Binder et al., 2007). However, catabolism of these amino acids, which provide the initial building blocks for the production of the acyl-CoAs used in bitter acid synthesis, is physically separated in cells from BCAA synthesis.

BCAA catabolism in plants is localized to mitochondria and peroxisomes (Binder et al., 2007).

Both the anabolic and catabolic pathways of BCAA synthesis and degradation are well studied

and a detailed understanding of both has emerged over the past two decades. Understanding of the biochemical pathways and gene regulatory networks involved in alpha acid production in hop, coupled with new DNA-marker and high through-put sequencing applications, has given breeders a new set of tools and greatly expanded genomic marker resources for future marker- assisted plant selection strategies.

An example of emerging discoveries in plant and animal gene regulatory networks, biochemical pathways, and genome analysis that can lead to new sets of DNA markers to screen for promising new cultivars is illustrated by a recent transcriptome study of alpha acid biosynthesis in hop (Clark et al., 2013). This study performed RNA sequencing (RNAseq) using high through- put sequencing techniques to analyze the transcriptomes of isolated lupulin glands, cones with glands removed, and leaves from high alpha acid hop cultivars. RNAseq is a type of transcriptome analysis sometimes referred to as whole transcriptome shotgun sequencing

(WTSS). RNAseq uses all the coding and non-coding RNA transcripts from a total RNA sample depleted of ribosomal RNA as the input RNA for high through-put sequencing. High through-put sequencing of this type of RNA sample provides the sequence and quantity all RNA transcripts sequenced. RNAseq datasets obtained by (Clark et al., 2013) were analyzed for RNA transcripts related to both catabolic and anabolic precursor pathways of BCAAs. When combined with the current understanding of BCAAs metabolic pathways the authors were able to identify and quantify genes involved in alpha acid biosynthesis. An analysis of this type would be impossible to perform without the extensive knowledge of the biochemical pathways, gene regulatory

12 elements associated with gene expression used in alpha acid biosynthesis, and high through- put sequencing techniques to which the authors had access. As mentioned above, the biosynthesis of alpha acids found in hop requires the synthesis and degradation of BCAAs.

BCAA biosynthesis from pyruvate in plants involves eight enzymes and several precursors. A distinctive attribute of BCAA biosynthesis is that valine and isoleucine are produced via parallel pathways. However, both use identical enzymes (Figure 6). Branched-chain aminotransferase

(BCAT) is used in the last anabolic step of BCAA synthesis and in the first step of BCAA catabolism to create the acyl-CoAs used in alpha acid synthesis. The anabolic and catabolic pathways are physically separated. BCAT structure and function have been studied in tomato and arabidopsis. Seven isoforms have been identified in tomato and six in arabidopsis (Diebold et al., 2002, Maloney et al., 2010, Kochevenko et al., 2012).

Results from Clark et al., (2013) identified four possible BCAT gene sequences in the hop genome. Two, denoted Humulus lupulus BCAT1 (HlBCAT1) and HlBCAT2 were plentiful in lupulin glands, with HlBCAT1 showing specificity to lupulin tissue. Both of these gene transcripts were confirmed by reverse transcription quantitative real-time PCR (RT-qPCR). As predicted by the RNAseq data, HlBCAT1 was confined to lupulin tissue and HlBCAT2 was found to be transcribed in both leaf and lupulin tissue. Alignment between the two genes shows 70% identity. Kinetic assays using HlBCAT1 and HlBCAT2 expressed in and purified from E. coli showed both enzymes were able to perform the anabolic and catabolic functions (Clark et al.,

2013). This contrasted with earlier work with tomato BCATs that indicated each BCAT was

13 functional in either the catabolic or anabolic direction, but not both (Maloney et al., 2010).

Using a method described by Emanuelsson et al.(2000) that forecasts protein subcellular localization based on N-terminal amino acid sequence, it was predicted that HlBCAT1 would localize to mitochondria and that HlBCAT2 would be directed to plastids.

Figure 6. BCAA biosynthesis from pyruvate in plants. Valine and isoleucine are produced via parallel pathways; both use identical enzymes. Branched-chain aminotransferase (BCAT) is used in the last anabolic step of BCAA synthesis and is the first step of BCAA catabolism to create the acyl-CoAs used in bitter acid synthesis. (Adapted from Maloney et al., 2010.)

Phylo genetic analysis using protein sequences of BCATs from hop, tomato, and other dicots showed a separation into two clades; anabolic/plastid and catabolic/mitochondrial. HlBCAT1 clustered into the catabolic/mitochondrial clade and HlBCAT2 clustered into the anabolic/plastid clade. This clustering is in agreement with the predicted subcellular localization based on N-terminal amino acid sequences. HlBCAT4, which lacks a signal

sequence, was clustered with BCATs that showed no apparent N-terminal localization signal.

In addition to the BCATs described above, Clark et al., (2013) identified several other genes that were differentially transcribed in the hop tissues examined. Some of these such as fructokinase, neutral alkaline/invertase and sucrose synthase are important in sucrose metabolism, and were upregulated in lupulin versus leaf tissue. This is consistent with the high energy requirements for the synthesis of the secondary metabolites produced in lupulin glands of the high acid content producing hops examined. Transcripts of genes from the methyl-D- erythritol 4-phosphate (MEP) pathway were also shown to be upregulated in lupulin gland tissue versus leaf tissue. This also could be expected as MEP pathway metabolites are essential for bittering acid prenylation (Binder et al., 2007). Several other gene transcripts reported by

Clark et al., (2013) to be more abundant in lupulin vs leaf or cone tissue, with glands removed were also directly involved in the BCAA/bittering acid pathways. The most abundant transcript in the lupulin gland dataset was VPS, which is involved in the final steps of bitter acid synthesis

(Figure 4).

In total, Clark et al. (2013) collected eleven RNA samples from tissues of four high alpha acid producing cultivars: ‘Nugget’, ‘Taurus’, ‘Magnum’, and ‘Apollo’. Tissue samples included were: lupulin gland composites, cones with glands removed, and leaves. cDNAs from each of these samples produced 15.7 to 24.7 million RNAseq reads that were assembled into approximately

170,000 contigs, with an average length of 745 bp. Among these contigs were 48 transcription factors that were greatly upregulated in lupulin tissue vs leaf or cone tissue void of glands. A

15 subset of 11 of these transcription factors had a greater than ten-fold increase, ranging from

10.1 to 498.2. A detailed discussion of gene regulatory networks and the regulatory elements involved in gene regulation, including the function of transcription factors is given in latter sections of this thesis.

The extensive transcriptome data reported by Clark et al. (2013) using high through-put sequencing technology significantly expanded lupulin gland transcriptome expressed sequence tags (EST) obtained by automated Sanger sequencing (Nagel et al., 2008; Wang et al., 2008; Wang and Dixon, 2009). These types of large scale transcriptome analysis, when combined with current knowledge of the biochemical pathways and gene regulatory elements associated with gene expression, are capable of providing new understandings of the genes connected with the production of primary and secondary metabolites associated with all aspects of plant growth, development, and disease resistance.

Large, genome-wide RNAseq experiments that use high through-put sequencing and smaller scale transcriptome analysis of coding and non-coding RNAs using ESTs, microarrays, and targeted northern blot techniques are not restricted to the analysis of metabolites produced by healthy plants and plant tissues, but have also been used to study changes in gene expression between diseased and disease-free plants and plant tissues. Several gene products associated with the molecular mechanisms and gene regulatory networks that govern a plants' tolerance or sensitivity to specific pathogens have been revealed. Genes identified by such studies, like the metabolite studies described above, provide genetic markers that can be used by breeders to identify plants that have a genetic profile that forecasts the probability of a plant to be tolerant or sensitive to specific or related plant pathogens such as disease causing fungi, bacteria, viruses, or viroids.

Viroids.

Viroids were first described by Diener (1971) as the smallest infectious agent of plants. They consist of extensively structured, circular, single stranded, RNA molecules ranging in size from

246 to 401 nucleotides, and lack a protective protein coat or other associated proteins. In contrast to all other plant pathogens such as fungi, bacteria, and viruses, no viroid has been shown to code for a protein or peptide product (Kovalskaya and Hammond, 2014). However, despite this lack of protein coding, viroids are capable of parasitizing plants and using an infected cells' transcriptional machinery for autonomous replication (Diener, 1974).

Interestingly, viroids seem to be constrained only to the plant kingdom, as no disease in animals has ever been associated with a viroid. Some studies have made comparisons between the

Hepatitis delta virus (HDV) and viroids. Like viroids the HDV genome is a highly structured, single stranded, circular RNA molecule. However, unlike viroids, its genome codes for a protein, the Hepatitis D antigen (HDAg), and, at approximately 1700 nucleotides, it is four times larger than the largest known viroid (Makino et al., 1987). An additional dissimilarity between viroids and HDV is the inability to replicate autonomously or be transmitted independently. For both these events, HDV depends on the Hepatitis B virus (HBV), and as such HDV, is considered a satellite of HBV (Elena et al., 1991).

The first recognized viroid to be characterized was Potato spindle tuber viroid (PSTVd) (Diener,

1971; 1974). However, potato spindle tuber disease, which is caused by PSTVd, was first reported in Solanum tuberosum L. by W.H. Martin in 1922. Symptoms of infected plants

17 included overall stunting of plants, and lengthened abnormally-shaped potatoes. Since Dieners' explanation of the cause of potato spindle tuber disease, several plant diseases worldwide in monocots, dicots, herbaceous, and woody plants have been shown to be the result of viroid infection. Examples include: tomato chlorotic dwarf, coconut cadang-cadang, citrus bark cracking, apple dimple fruit, avocado sun blotch, peach latent mosaic, and hop stunt. Viroids are assigned to one of two families; Pospiviroidae or Avsunviroidae (table 1).

Length Species (nt) Known hosts Family Pospiviroidae Pospiviroid Potato spindle tuber (PSTVd) 341-364 potato, tomato, avocado Chrysanthemum stunt (CSVd) 348-356 chrysanthemum

Citrus exocortis (CEVd) 366-475 citrus, tomato

Tomato apical stunt (TASVd) 360-363 tomato

Tomato chlorotic dwarf (TCDVd) 360 tomato

Hostuviroid Hop stunt (HSVd) 294-303 hop, cucumber, citrus, grapevine, Prunus spp. Cocadviroid Coconut cadang-cadang (CCCVd) 246-301 coconut palm

Citrus bark cracking (CBCVd) 284-286 citrus

Hop latent (HLVd) 255-256 hop

Apscaviroid Apple scar skin (ASSVd) 329-333 apple, pear

Apple dimple fruit (ADFVd) 306 apple

Apple fruit crinkle (AFCVd)z 368-372 apple

Citrus bent leaf (CBLVd) 315-329 citrus

Family Avsunviroidae Avsunviroid Avocado sun blotch (ASBVd) 239-251 avocado

Pelamoviroid Chrysanthemum chlorotic mottle (CChMVd) 397-401 chrysanthemum Peach latent mosaic (PLMVd) 335-351 peach, nectarine

Table 1. Representative sample of viroids, their known host and the associated viroid family Pospiviroidae or Avsunviroidae

Principal differences that determine viroid classification are: subcellular location of replication, existences of hammerhead ribozyme motifs, and types of secondary structure organization

(Kovalskaya and Hammond, 2014). Viroids of the family Avsunviroidae replicate in plastids, primarily in chloroplast, by producing longer than unit length oligomers that are processed to unit length by autolytic cleavage activity of internal hammerhead ribozyme moieties (Hutchins et al., 1986).

Pospiviroidae viroids replicate in the nucleus with the aid of host specific DNA-dependent RNA polymerase II (Mühlbach and Sänger, 1979) which produces longer than unit length oligomers similar to Avsunviroidae viroids. However, Pospiviroidae family members lack hammerhead ribozyme moieties, and rely on host coded enzymes for cleavage to unit length RNA molecules

(Branch et al., 1988). Viroids from both families are single-stranded circular RNA molecules that, via intramolecular base pairing, are partially double-stranded with secondary structures that are highly resistant to thermal degradation. Secondary structures include: helices, hairpins, pseudoknots, and loops. Each of these structures, depending on the viroid, have been shown to be important in one or more viroid life cycle function such as replication, intracellular or systemic transport, and cytopathology. Structural analysis of several Avsunviroidae viroids revealed a secondary structure that is typically a complex multibranched arrangement of hairpins, pseudoknots, and loops that form hammerhead ribozymes, which are unique to

Avsunviroidae (Flores et al., 1999) (Figure 7A). Interestingly, Avocado sun blotch viroid, the type species of Avsunviroidae lacks much of the multibranched character common to other members of this family (Symons, 1981).

Unlike members of Avsunviroidae, members of Pospiviroidae have a simpler, less complex secondary structural arrangement (Figure 7B). The most striking feature of Pospiviroidae viroids is their rod-like structure (Riesner et al., 1979) which contains five sequence-related domains: terminal left (TL), terminal right (TR), central conserved region (CCR), pathogenicity (P), and variable (V) domains (Keese and Symons, 1985). The CCR is shaped by two segments of

20 conserved nucleotides in the upper and lower strands flanked by a faulty inverted repeat

(McInnes and Symons, 1991). The TL domain is characterized by two mutually exclusive motifs; either a terminal conserved hairpin (TCH) or a terminal conserved region (TCR) (Flores et al.,

1997). Mutagenesis at the single nucleotide level has shown that each of these domains has one or more functional roles in cell-to-cell or systemic movement, replication, sub-cellular localization, and pathogenicity (Flores et al., 2012).

Figure 7. A) Multibranched secondary structure showing hairpins, pseudoknots, and loops that form hammerhead ribozymes, which are unique to Avsunviroidae viroids. B) Less complex secondary rod like structure of Pospiviroidae viroids and their five sequence related domains; terminal left (TL), terminal right (TR), highlighted central conserved region (CCR), pathogenicity (P), and variable (V) domains. [A Adapted from Diener, T. O. (2003) ; B) from Wikimedia Commons, the free media repository https://commons.wikimedia.org.]

Key aspects of viroid pathogenicity is their ability to colonize host cells, move to, or be localized to the site of replication, and then move systemically through the plant’s vascular system, primarily the phloem tissue, in a source to sink direction. Because viroids do not code for proteins, each of these steps depends on host encoded RNA-binding proteins. It has been shown that a dimeric lectin from cucumber phloem exudate, phloem protein 2 (PP2), is capable of interacting with Hop stunt viroid (HSVd) RNA, and may facilitate the systemic movement in plants of HSVd, and potentially other viroids (Owens et al., 2001). Other proteins, such as

Virp1, from tomato, which has an atypical RNA binding site and a nuclear localization motif was shown to interact with HSVd and PSTVd (Maniataki et al., 2003). HSVd has a lower affinity for

Virp1 than PSTVd, and this may be a possible reason for the lower infectivity rate of HSVd in tomatoes compared to that of PSTVd (Maniataki et al., 2003). Further analysis of the TR hairpin loop showed that minor variations in the 12 bp loop were likely responsible for the different affinities. Such sequence variations between members of a viroid family are not unexpected; however, minor variations have also been observed between closely related but non-identical isolates of the same viroid. Such variations result from the high mutation rate associated with viroid replication, and can affect both expression of symptoms and pathogenicity (Domingo et al., 1996; Flores et al., 2006). There are many examples of viroids that show sequence variations among isolates, but further description of these changes and their effects are restricted to HSVd.

HSVd is a member of the Pospiviroidae family, like other members of this family, HSVd is a single-stranded, circular RNA molecule with a rod-like secondary structure. Several natural occurring variants of HSVd that cause disease in a wide host range have been isolated and sequenced; hosts include hop, grapevine, citrus, cucumber, and stone fruits (Ohno et al., 1983;

Sano et al., 1984, 1985, 1986; Kawaguchi-Ito et al., 2009). Isolates range in size from 294 to 303 bp, and have between 1 and 16 nucleotide variations.

In a 15-year evolutionary study, Kawaguchi-Ito et al. (2009) showed that a single hop cultivar,

Humulus lupulus L. cv. Kirin II, infected with HSVd isolates from grapevine (HSVd-grapevine), plum (HSVd-plum), citrus (HSVd-citrus) and hop (HSVd-hop) had differing times of symptom onset, with varying degrees of symptom severity and expression patterns. Plants infected with

HSVd-grapevine and HSVd-hop started to show mild stunting symptoms in year four, and in year five the stunting symptoms became more pronounced. By year seven, the stunting was severe. Additional 'stunt disease' symptom leaf epinasty appeared in year seven. Plants infected with HSVd-citrus showed severe vine stunting and epinasty in year one. By year two, the lateral branches drooped severely and vine diameter was significantly reduced. Symptom severities caused by HSVd-plum were judged to be between those of HSVd-hop/grape vine and

HSVd-citrus. Alpha acid levels as measured by percent dry cone weight decreased significantly in all infected plants by year two, ranging from 46% to 55% reduction in plants infected with

HSVd-hop, -grapevine, or -citrus and to 25% in plants infected with HSVd-plum. By year three the reduction increased to 50% to 60% in HSVd-hop, -grapevine, and -citrus plants and ranged from 28% to 45% in HSVd-plum infected plants.

In addition, Kawaguchi-Ito et al. (2009) monitored the sequence of each HSVd strain for change throughout the 15-year study. Results from the cataloged sequence data showed that HSVd- grapevine, while replicating in hop for extended periods of time, produced several combinations of mutations among five nucleotides that evolved into the major HSVd-hop viroid strains found in hop gardens throughout the world. Based on this evidence and the geographical distribution of hop stunt viroid disease prior to its worldwide spread, the authors suggest that HSVd-grapevine viroids were the initial inoculum of hops, but mutated into HSVd- hop strains, which eventually became the primary source of inoculum for the spread of HSVd- hop.

In a variation of the above study (Yaguchi and Takahashi, 1984) used the same HSVd strain to infect different plant hosts. That study investigated the relationship of HSVd and symptom expression in eight different cucurbitaceous species including 26 cucumber cultivars. All 26 cucumber cultivars developed symptoms as a result of HSVd infection, but the severity of symptom expression ranged from mild to severe in a cultivar-dependent fashion. Severe symptoms presented as shortened stem length, small rugose leaves with extensive epinasty, and reduced flora development. Intermediate symptom expressing plants showed similar leaf and flora development as severe expressing plants; however, stem shortening was not as pronounced. Mild expressing plants showed a slight stunting in mature plants, with normal floral development; early leaf rugoses developed, but diminished as the plant matured.

Severe symptom expression patterns in the cucumber Cucumis sativus cv. Suyo infected with

HSVd were shown by Kojima et al., (1983) to be associated with abnormal cell wall development that had a ribbed profile and asymmetrical thickness in epidermal, palisade parenchyma, spongy parenchyma and vascular tissue cells. These abnormalities appeared to be the cause of the structural deviations associated with overall cell shape that leads to the resulting external symptoms observed by Yaguchi and Takahashi (1984). Another modification observed in HSVd infected cucumber leaves was chloroplast disintegration, breakdown of tonoplast, and accumulation of granules within the cell. Momma and Takahashi (1982) observed similar cytological abnormalities in the HSVd-infected hop Humulus lupulus L. cv.

Shinshuwase. HSVd-infected tissue had abnormal cell wall development similar to HSVd infected cucumber including ribbed, asymmetrical thickening of cell walls. Also, chloroplast thylakoid membranes were disorganized with poor stacking of grana. One additional cytopathic defect noted by Momma and Takahashi (1982) in HSVd-infected hop that did not appear to occur in cucumber was the presence of a malformed cuticle layer. In HSVd-free tissue, the cuticle layer appears to have wax deposits arranged in a folding pattern covering each epidermal cell. In HSVd-infected tissue, the wax deposits are missing. These cytological abnormalities are most likely the result of a combination of interference between one or more biochemical pathways, and explicit interference of one or more normal gene regulatory pathways for specific host genes. Qi and Ding (2003) were able to associate diminished cell wall expansion, which lead to stunting in PSTVd infected tomato, to the down-regulation of the

LeExp2 gene.

LeExp2 codes for expansin, a nonenzymatic protein found in cell walls and important in several cell growth functions.

A recent comprehensive study by Owens et al., (2012) of gene expression patterns in sensitive and tolerant cultivars of tomato plants free of and infected with PSTVd showed that more than half of the 10,000 genes arrayed on the Affymetrix Tomato GeneChip had altered gene expression patterns for the cultivar ‘Rutgers’ infected with PSTVd. A second cultivar,

‘Moneymaker’, which has reduced symptoms expression relative to ‘Rutgers’, had similar results. In addition, a transgenic ‘Moneymaker’ expressing small interfering RNAs (siRNAs) coded from PSTVd showed fewer differences, but still had extensive changes in its gene expression patterns. To validate the microarray results the authors choose six genes for quantitative reverse transcription-polymerase chain reaction (qRT-PCR); results from the qRT-

PCR were equivalent to that of the microarray. These results showed that disease severity was a poor predictor of the number of genes differentially expressed and that the pattern or direction of the changes, up or down regulated, were similar for each plant.

Differentially expressed genes were categorized into one of three groups: cellular component, biological process, and molecular function (Owens et al., 2012). Using these criteria, it was evident that genes involved in chloroplast function were down regulated in all PSTVd-infected plants. Genes coding for proteins linked with the nucleus, plasma membrane, cell wall, ribosomes, and apoplast were up-regulated in PSTVd-infected ‘Rutgers’, but not in PSTVd-

26 infected ‘Moneymaker’. Nineteen genes involved in hormone anabolic and catabolic processes showed changes in gene expression patterns. These included genes in the gibberellin (GA), jasmonic (JA) and abscissic acid (ABA), brassinosteroid (BR), cytokinin (CK), and ethylene (ET) biosynthetic pathways. JA and ET are known to have prominent roles in directing a plants response to biotic and abiotic stress. Three proteins known to be important in JA signaling pathway are coronatine insensitive 1 (COI1), jasmonate insensitive 1 (JIN1/MYC2), and jasmonate resistant 1 (JAR1). JAR1 codes for a synthetase involved in the conversion of isoleucine to JA, and was shown to be down-regulated in both the infected and transgenic

‘Moneymaker’ plants. Neither COI1 nor JIN1 showed changes in their gene expression patterns. Additionally, several genes important to the ET signaling pathway showed changes in their gene expression patterns. A member of the ET receptor subfamily 2 (EIN4), a component of a transcriptional cascade important in the biosynthesis of several transcription factors and two EIN3 dependent transcription factors (ERF1 and ERF2) were up-regulated in PSTVd-infected plants. Similar changes in the gene expression patterns for components of the GA, ABA, BR, and CK signaling cascades were observed.

A detailed understanding of all the components involved in gene regulation are an important aspect of any large scale analysis of differential gene expression patterns. In addition, specific knowledge of individual gene regulatory pathways involved in anabolic and catabolic pathways of primary and secondary metabolites involved in plant growth, development, and response to abiotic and biotic stress is important as well. Whatever the mechanisms used by viroids to

27 cause symptoms in plants, it will have some gene regulatory component; most likely it will be a combination between a metabolic pathway disturbance and interference with one or more gene regulatory networks.

Operons.

Pardee, Jacob, and Monod (Pardee et al., 1959) were the first to describe a gene regulatory mechanism linked to the expression of a protein: the enzyme -galactocidase used by the prokaryote Escherichia coli (E. coli) in the metabolism of -galactosides. Over the last several decades, researchers have continued to build upon this genetic switch model first proposed by

(Pardee et al., 1959). As a result, it has become clear that the mechanisms that control gene regulatory events in both prokaryotes and eukaryotes are much more complex than that described even a decade ago.

Jacob and Monod (Jacob et al., 1960) used the term operon to describe a “group of genes with expression coordinated by an operator”. In this model, a continuous stretch of DNA containing a group of genes is under the control of one promoter and one operator (Jacob and Monod,

1961). Promoters lie upstream of the transcriptional start site (TSS) and have unique DNA sequences that are recognized by DNA-dependent RNA polymerases. One of the first steps in transcription is binding of the RNA polymerases to a promoter. Operators, on the other hand, are unique sequences of DNA recognized by repressors. They are located in close proximity to promoters. The binding of repressors to an operator creates a physical barrier to the binding of

RNA polymerases, and thus represses transcription.

Operons in prokaryotes typically involve a cluster of genes regulated by only one promoter and one operator (Salgado et al., 2000). A gene cluster is defined as “a set of two or more non- homologous genes encoding enzymes from the same pathway (to be distinguished from the gene clusters resulting from tandem duplication and consisting of homologous genes)”

(Boycheva, 2014). For example, the three structural genes β-galactosidase, lactose permease, and galactoside O-acetyltransferase, responsible for the metabolic pathway involved in the transport and catabolic break down of the -galactoside lactose in E. coli, are considered a gene cluster and are regulated by a single promoter and operator (Jacob and Monod, 1961). In prokaryotes structural genes in gene clusters are typically transcribed as a single polycistronic transcript (Jacob et al., 1960). Gene clusters in eukaryotes that are controlled by a shared regulatory promoter or promoters and operators are most often transcribed independently, each as an individual messenger RNA rather than as a polycistronic transcript (Boycheva, 2014).

However, there are examples of eukaryotes that display types of polycistronic processing that can be divided into two generalized methods. The first makes use of internal ribosome entry sites (IRES), or some other form of ribosomal reentry following a stop codon. The second involves alternative splicing events to create multiple monocistronic mRNAs from a single, larger polycistronic pre-mRNA transcript (Blumenthal, 2004).

Frey et al. (1997) revealed one of the first gene clusters in plants; this cluster involved five genes Bx1 through Bx5. The products from these five genes are involved in the production of a secondary metabolite, 2,4-dihydroxy-1,4-benzoxazin-3-one (DIBOA), an important element in

29 the detoxification of herbicides and pesticides, and in protecting host plants (Poaceae family) from a diverse cross section of insects, fungi and bacteria plant pathogens (Niemeyer, 1988).

Additional examples of gene clusters in plants include the triterpene biosynthetic gene clusters in oat (Qi et al., 2004), and the momilactone gene cluster in rice (Shimura et al., 2007, Sue et al.,

2011). Each of the genes in these clusters and in other plant gene clusters discovered to date code for enzymes used in the production of secondary metabolites. No operon gene clusters or

“operon-like gene clusters” have been described for primary metabolites in plants (Boycheva,

2014). However, all genes, whether part of a gene cluster or a single gene regulated independently of other genes by a unique regulatory sequence, require a RNA polymerase

(RNAP) recognition sequence within a core promoter (CP), several coactivators and multiple transcription factors bound to the RNAP, and a promoter as part of the basal transcriptional machinery (Juven-Gershon and Kadonaga, 2010; Kumari and Ware, 2013; Boycheva, 2014).

Coactivators and transcription factors are distinguished from each other based on the ability to bind a DNA recognition site or not. Transcription factors are capable of binding DNA and coactivators are not; instead, coactivators bind to other proteins in the transcription complex.

Gene regulation

Genes that code for proteins, microRNAs (miRNA), and other non-coding RNAs are transcribed by the DNA-dependent RNA polymerase II (pol II), which binds as a holoenzyme along with different transcription factors and coactivators to a promoter sequence close to the transcription start site (TSS) of the gene being transcribed. Promoter sequences can be divided

30 into two regions; proximal and distal. With the TSS designated as the +1 nucleotide position, the proximal region is generally between the -40 and + 40 nucleotide positions. This region encompasses the core promoter element (CPE) which is responsible for binding pol II (Juven-

Gershon and Kadonaga, 2010). Distal regions contain cis-regulatory elements which work in combination with the CPE to regulate gene transcription (Kumari and Ware, 2013). Additional regulatory elements include enhancer sequences which, in animals, can lay as much as a million base pairs (bp) upstream or downstream of the gene they help control (Pennacchio et al.,

2013). Plant enhancers however, are generally within 1000 bp of the TSS for the gene they regulate (Komarnytsky and Borisjuk, 2003).

Core promoter elements in plant and animal promoters contain the principal recognition sequence used to bind pol II during the formation of a pol II pre-initiation complex (PIC).

Interestingly, pol II, an enzyme composed of several subunits when purified, is incapable of recognizing or binding to a DNA recognition sequence without the aid of additional general transcription factors (GTFs) (Juven-Gershon and Kadonaga, 2010; Murakami et al., 2013). Some

GTFs involved in the formation of the pol II holoenzyme include transcription factors for RNA polymerase II A (TFIIA), TFIIB, TFIID, TFIIE, TFIIF, and TFIIH. GTFs comprise several functions: recruitment of DNA, unwinding of the DNA duplex, RNA synthesis of the first 25 bases, and formation of a stable RNA elongation complex (Murakami et al, 2013). Although the formation of every PIC requires a pol II recognition sequence and the use of several GTFs, the combination of GTFs and the pol II recognition sequence is not the same for every core promoter.

Additionally a core promoter can contain two or more CPEs (Burley, 1996).

Recently, core promoters have been categorized as focused or dispersed. This distinction is based on the number of nucleotides that can be used within the CP for transcription initiation.

Focused initiation CPs have tightly focused TSS over a very narrow range of nucleotides and dispersed initiation CPs utilize multiple weak TSS along a stretch of 50 to 100 nucleotides. Each of these; focused or dispersed CPs appear to utilize different GTF combinations and have different CPEs that are recognized by a pol II holoenzyme to form the PIC (Juven-Gershon and

Kadonaga, 2010).

Functional genomic sequence motifs in plant promoters or CPEs implicated in PIC formation have not been as extensively cataloged or studied as those in yeast and animals. One of the first recognized CPE genomic motifs, and the most studied in plants and animals, is the TATA- box motif (Breathnach and Chambon, 1981; Juven-Gershon and Kadonaga, 2010; Kumari and

Ware, 2013). Analysis of 79 TATA-box motifs in plant promoters identified that

TCACTATATATAG was the consensus sequence (Joshi, 1987). This motif is recognized by the

TATA box binding protein (TBP) subunit of the TFIID general transcription factor (Burley, 1996).

An analysis of arabidopsis core promoters by Molina and Grotewold (2005) found that approximately 29% of all arabidopsis promoters contain TATA-box motifs, centered at nucleotide position -32 relative to the TSS. A detailed structural analysis by Murakami et al.

(2013) of the yeast Saccharomyces cerevisiae PIC bound to the histidine 4 gene promoter, which contains a TATA-box motif, revealed that a complex of 32 proteins, including pol II, were

32 arranged into two distinct regions, which together formed the PIC. Other PIC complexes that recognize a TATA-box motif have been described in other biological systems. TATA-box promoters account for approximately 19% of rice promoters (Civan and Svec, 2009), 13% in yeast and 10% in humans (Yang et al., 2007).

Promoters of genes that are “TATA-less” have been described. They are often expressed constitutively and are involved primarily in regulating housekeeping genes (Basehoar et al.,

2004; Yang et al., 2007). In contrast, TATA-box genes studied in human and yeast genomes linked with tissue specificity, are often expressed as a response to stress, and are less widespread than “TATA-less promoters” (Kumari and Ware, 2013). CPEs associated with

“TATA-less promoters” include the X core promoter element 1 (XCPE1), upstream TFIIB recognition element (BREu), initiator (Inr), downstream promoter element (DPE), motif ten element (MTE), and downstream TFIIB recognition element (BREd) (Juven-Gershon and

Kadonaga, 2010). As mentioned above, a single promoter may contain two or more CPEs.

Deng and Roberts (2005) showed that the TATA-box and (BREd) CPEs were combined into a single promoter, and Hsu et al. (2008) showed a regulatory network in cultured drosophila cells that involved crosstalk between TATA-box and DPE motifs located in a single promoter. In this gene regulatory network, the TATA-box binding protein (TBP) subunit of TFIID binds and activates TATA-dependent transcription, and at the same time, down regulates DPE-dependent transcription. In contrast Mot1 and NC2, two functionally related proteins know to disrupt TBP binding to the TATA-box repressed TATA dependent transcription and up regulated DPE- dependent transcription.

To date, many of the functional CPEs (XCPE1, BREu, DPE, MTE, and BREd) associated with yeast and animal promoters have not been described in plant regulatory networks (Schmidt and

Bancroft, 2011). Many of the general transcription factors, e.g., TFIID, TFIIB, and others involved in PIC formation described above, are common to plants, animals, and yeast. In addition, spatial relationships between the TSS and TATA-box motif in plants and animals are well conserved (Yoshiharu et al., 2007). However, reported motifs surrounding the TSS of plants and animals have differences (Fujimori et al., 2005). For example, the initiator (Inr) CPE in plants and animals has a different size and consensus sequence. In Arabidopsis thaliana and Oryza sativa, a region described as the Y patch, is rich in pyrimidines located between positions -10 to

-60 bp and centered at bp -13 but is absent in yeast and animal promoters (Yoshiharu et al.,

2007). Another putative CPE, found in arabidopsis that has no homologue in animals is the GA element. This CPE discovered by in silico analyses is in close proximity of the TSS and can be found up stream or downstream from the Inr CPE (Yamamoto et al., 2009). Unlike the Y patch that is generally coupled with the TATA-box motif, the GA element excludes the TATA-box CPE because its relative position within promoters overlaps that of the TATA-box. Though the GA element has little sequence homology to animal CPEs, it does have functional characteristics similar to animal CpG islands, (Yamamoto et al., 2009). It was estimated that 21% of arabidopsis promoters that control structural genes are associated with GA elements (Schmidt and

Bancroft, 2011).

Core promoter elements such as the GA element, Y patch and TATA-box motif are all located in the proximal promoter region within 40 to 50 bp of the TSS. Other important regulatory elements which are located in distal regions of promoters are the cis-regulatory elements (CRE).

These non-coding DNA sequences are recognized by a broad array of trans-acting transcription factors (TF), which when bound to a CRE can modulate transcriptional activity by enhancing or repressing the function and or formation of the PIC described above (Ong and Corces, 2011).

Several TFs and their CRE targets have been identified in animals and plants. Recently, Yant et al. (2010) reported a genome-wide mapping study that revealed the mapping profile of the

Arabidopsis thaliana AP2 transcription factor to multiple CREs in the arabidopsis genome. The

AP2 transcription factor is known to have several functions in a variety of plant development processes such as seed maturation, stem cell continuation, and floral development (Ohto et al.,

2001, 2005). The molecular mechanisms and gene regulatory networks by which this TF is able to modulate gene expression is illustrative of the added complexity of gene regulatory events that involve CREs and their related TFs.

AP2 and five redundant TFs (TOE1, TOE2, TOE3, SMZ, and SNZ), which contain an AP2 binding domain, were shown to modulate flower development by binding to and down regulating transcription from promoters in the; SUPPRESSOR OF OVEREXPRESSION OF CONSTANS1 (SOC1) and AGAMOUS (AG) loci, and by up regulating expression of a floral repressor AGAMOUS-

LIKE15 at a third loci (Yant et al., 2010). Additionally, AP2 and its five redundant TFs were shown to bind to two microRNA genes (miR156 and miRNA172) that control AP2 expression.

The binding of AP2 to these two genes has opposing affects: AP2 binding up regulates miRNA156, and down regulates miRNA172. Because each of these microRNAs positively modulates AP2 expression, the overall effect is creation of a negative feedback loop that controls AP2 expression (Yant et al., 2010).

The AP2 binding regions in SOC1 and AG loci are dissimilar. The AP2 binding region for SOC1 is located at the 5’ end, 1.6 kb from the TSS. A second floral development gene SEPALLATA3

(SEP3) was also shown to contain an AP2 binding region in the same location as SOC1. In contrast, the AP2 binding region for AG was located at the 3’ end of the 2nd intron (Yant et al.,

2010). Multiple AP2 binding sites are also present in AP2s’ own locus and in the five redundant

TF genes listed above. The number and relative position to the TSS position of each of these

CREs varied for each TF. These arrangements highlight the fact that CREs are mobile in relation to the TSS of the genes they modulate and that multiple cis-regulatory elements, for the same or different TFs, are utilized to regulate gene transcription.

Small RNAs.

In contrast to TFs, small RNAs (sRNAs) in plants are capable of directly regulating transcription and translation at several points along the gene to transcript to protein pathway (Voinnet,

2009). Points of regulatory control include transcriptional gene silencing (TGS), chromatin remodeling, posttranscriptional gene silencing (PTGS) transcript cleavage, accelerated transcript decay, and translation inhibition (Eulalio et al., 2008). Each of these regulatory

36 mechanisms is generally specific to a class of sRNA, but overlapping functionality between sRNAs does exist (Voinnet, 2009).

The most abundant sRNAs in plants are the cis-acting short interfering RNAs (siRNAs) (Axtell,

2013). These siRNAs are often referred to as heterochromatic siRNAs because they are involved in chromatin remodeling via methylation and acetylation of histones and cytosine residues in the loci that code for them (Matzke et al., 2009). The majority of heterochromatic siRNAs have a length of 23 to 24 nucleotides, which is longer than other plant sRNAs, that are typically 20 to22 nt long (Ruiz-Ferrer and Voinnet 2009). In addition, heterochromatic siRNAs are produced from a double-stranded precursor RNA by RNA dependent RNA polymerase

(RDR2) and a Dicer-like protein (DCL3). Furthermore, heterochromatic siRNAs depend on a specific subset of ARGONAUTE proteins (AGO4 or AGO6) for their RNA-directed DNA methylation (RdDM) function (Voinnet, 2009; Havecker et al., 2010).

Heterochromatic siRNAs or cis-acting siRNAs are only one of several types of siRNAs produced from double stranded RNA (dsRNA) precursors. Other siRNAs derived from a double stranded precursor include hairpin RNA (hpRNA) that contain inverted repeats. Expression of transgenes containing inverted repeats established in plant cells are used to produce hpRNAs that can be processed into functional siRNAs that are capable of PTGS. This strategy, known as IR posttranscriptional gene silencing (IR-PTGS), has been used to down-regulate endogenous mRNAs in plants (Smith et al., 2000; Wroblewski et al., 2014).

Additional siRNAs are derived from dsRNA precursors are secondary siRNAs and natural antisense transcript siRNAs (nat-siRNAs) (Axtell, 2013). Nat-siRNAs, like hpRNAs, do not depend on an RDR for formation of a double stranded precursor. They rely on Watson-Crick base pairing between two separately transcribed, but complementary RNAs. Katiyar-Agarwal et al.

(2006) identified a nat-siRNA (nat-siRNA-ATGB2) that is induced in tomato by a specific pathovar of Pseudomonas syringae (Pst-avrRpt2) that codes for a specific bacterial effector avrRpt2. Effectors such as avrRpt2 are directly or indirectly recognized by host R proteins. R proteins are expressed from resistance genes (R-Genes) that when expressed by plants provide disease resistance against pathogens by counteracting avirulence (Avr) gene products, such as avrRpt2 which are capable of silencing or attenuating a plants immune response (Eulgem,

2006). The specific host R gene, RPS2, induced formation of nat-siRNA-ATGB2. This nat-siRNA in combination with the NDR1 host gene product prompted a RPS2-directed immune-like response in tomato plants infected with Pseudomonas syringae (Pst-avrRpt2). The natural antisense transcripts (NATs) required for the production of nat-siRNA-ATGB2 was determined to be an overlapping region between a GTP-binding protein gene (Rab2-like) and a pentatricopeptide repeat-like protein gene (PPRL). The mature 22nt nat-siRNA-ATGB2 sequence is complementary to a 3’ UTR region of PPRL. Infection of tomato plants with Pst-avrRpt2 resulted in production of nat-siRNA-ATGB2 and a significant reduction of PPRL transcripts.

Conversely, infection of tomato plants with a different P. syringae pathovar (Pst-avrRpm1), which lacks the avrRpt2 gene, showed no induction of nat-siRNA-ATGB2, and consequently no reduction of PPRL transcripts.

Similarly, Borsani et al. (2005) showed that salt tolerance in arabidopsis was, in part, controlled by two nat-siRNAs. The first of the two is produced via an overlapping region between the arabidopsis genes P5CDH and SRO5, is 24 nt in length, and is named 24-nt SRO5-P5CDH nat- siRNA. The second is produced from cleavage products of P5CDH transcripts created using the

24-nt SRO5-P5CDH nat-siRNA to guide P5CDH cleavage. This second nat-siRNA is further processed by DICER-Like protein 1 (DCL1) to a 21 nt nat-siRNA termed 21-nt P5CDH. The production of both nat-siRNAs is dependent on the induction of the SRO5 gene as a result of salt stress. Production of 21-nt P5CDH nat-siRNA guides cleavage of P5CDH mRNA; down regulation of P5CDH leads to proline accumulation that is important for salt tolerance. Similar regulatory loops involving sRNAs of every type are being identified regularly (Voinnet, 2009;

Axtell, 2013).

Another distinct class of sRNA molecules is the secondary siRNAs. Secondary siRNAs develop from double stranded precursors produced from the activity of RDR6 on the cleavage products generated as a result of transcript cleavage guided by a different sRNA (Axtell, 2013). Mature secondary siRNAs are produced via processing of the double-stranded precursor by DCL 4.

Production of the initial cleaved transcript that results in the template used by RDR6 for the formation of a double-stranded secondary siRNA precursor is often by a miRNA or another secondary siRNA (Allen et al., 2005). Gomez et al. (2008) showed that RDR6 was important for development of symptoms in Nicotiana benthamiana plants infected with HSVd, independent of HSVd titer levels.

A subset of secondary siRNAs is the phased secondary siRNA molecules. Phased secondary siRNAs have the same bioactive parameters as other secondary siRNAs, but differ in their biogenesis from other secondary siRNAs. Phased secondary siRNAs are produced by the sequential cutting of a RNA molecule that was previously targeted by another sRNA for cleavage (Brodersen and Voinnet, 2006). As such, phased secondary siRNAs produce a unique pattern of sRNAs that cover several contiguous nucleotides within a single transcript. Phased secondary siRNAs are often associated with the down regulation of several genes within a gene family (Axtell, 2013). However, only a small portion of sRNA targeted silencing exchanges result in secondary siRNA production. Attributes of the first sRNA trigger, activity of the associated AGO, and characteristics of the initial RNA target help determine production of secondary siRNAs. Target RNA with abnormal 5’ or 3’ ends (Herr et al., 2006) and ones with two or more sRNA target sites or with near perfect complementary sRNA target sites have been shown to support production of secondary siRNAs (Axtell et al., 2006).

A fourth class of sRNA is the microRNAs (miRNAs). These sRNAs differ from the secondary siRNAs, nat-siRNAs and heterochromatic siRNAs discussed above in several ways. Unlike the above described sRNAs, which are derived from double-stranded RNA molecules, miRNAs are derived from single-stranded RNA molecules that form a hairpin loop, creating a secondary double-stranded rod-like structure held together by complementary base pairing. Thus, miRNA secondary structure has many similarities to hpRNA produced by transgenes containing inverted repeats, and to viroids of the Pospiviroidae family such as HSVd.

MicroRNAs are produced via DCL1 cleavage, and require a member from the AGO1- clade for their bioactivity. DCL1 cleavage of pre-miRNAs produces either a 21 or 24 nt miRNA, the latter are referred to as long miRNAs. In contrast to the 21 nt miRNAs the 24 nt long miRNAs have functional similarities to the heterochromatic siRNAs in that both remodel chromatin via acetylation and methylation events (Wu et al., 2010). However, 21 nt miRNAs regulate gene expression by directing the degradation of transcripts and repression of translation. No distinction has been observed that predicts which pathway (degradation of transcripts and repression of translation) will be used by any given 21 nt miRNA. The 21 nt miRNAs appear to be more evolutionarily conserved across the plant kingdom then the other sRNAs including the

24 nt long miRNAs (Cuperus et al., 2011).

Evolutionarily conserved sRNAs, including the miRNAs, have likely arisen through several gene duplication events and reordering actions (Axtell, 2013). Many of the evolutionarily conserved miRNAs regulate transcription factors that control fundamental gene regulatory networks that set the stage for early plant development events, such as organ specific cell fates and meristem development (Garcia, 2008). In contrast, evolutionarily recent miRNAs are regularly encoded by single-copy genes, are linage specific, often need induction by biotic or abiotic factors, and the proteins associated with their regulation cover effectively every facet of plant biology

(Voinnet, 2009).

Regardless of a sRNA’s evolutionarily past or the type of sRNA (miRNA, nat-siRNA, heterochromatic siRNA, or secondary siRNA), they have fundamental requirements for core enzymes important for their biogenesis and action. Three protein families essential to plant sRNA biogenesis and action are: RNA dependent RNA polymerases (RDR), Dicer-like enzymes

(DCL), and ARGONAUTE proteins (AGO). RDRs make a complementary second-strand RNA to a single-strand template, producing a dsRNA precursor. DCLs are endonucleases that cleave the dsRNA precursor releasing short, 20 to 24 nt long, dsRNA fragments (sRNAs) with 2 to 3 nt overhangs. AGO forms complexes with sRNAs, discarding one strand of the RNA duplex, forming an AGO/sRNA-loaded complex. The bound single-stranded small RNA fragment serves to specify the transcript target or DNA loci target via complementarity to the small RNA fragment. Positive target identification leads to one of several possible repressive activities coordinated by the linked AGO protein. Repressive activities include: transcriptional gene silencing (TGS) via chromatin remodeling, posttranscriptional gene silencing (PTGS) via transcript cleavage, accelerated transcript decay, and translation inhibition. As a result of these repressive activities, gene regulatory networks, and the proteins or non-coding RNAs for which they code are maintained at a steady state, down-regulated or up-regulated accordingly. Such regulation is important to every aspect of plant growth and development; including maintenance of healthy levels of primary and secondary metabolites and responses to abiotic and biotic stimuli, such as drought and pathogen related diseases.

Pathogenesis related proteins.

Plants have an extensive number of mechanisms to protect themselves against a wide variety of biotic and abiotic stresses such as wounding, extreme fluctuations in temperature and moisture, and pathogen attack. Plant reactions to these stresses are multifaceted, and involve the adjustment of gene regulatory networks and signal transduction cascades that control the expression patterns of primary and secondary metabolites, non-coding RNAs and protein- coding RNAs used by plant defense mechanisms (Muthamilarasan and Prasad, 2013). Abiotic stresses or invading pathogens induce physiological changes in plants such as cell wall thickening by callose deposition, and lignification to mitigate abiotic stress or prevent further pathogen invasion and/or damage (Ruiz et. al., 2011). These physiological changes can be associated with a localized hypersensitive response (HR) induced by invading pathogens. This response synchronizes an integrated set of metabolic reactions that facilitate rapid cell necrosis localized to cells proximal to the infection site (Hammond-Kosack and Jones. 1996). In addition, pathogen invasion can induce an innate plant immune response which involves expression of numerous unique proteins, collectively referred to as ``pathogenesis-related proteins” (PR)

(Spoel and Dong, 2012). Unlike proteins involved in localized cell necrosis and cell wall thickening, which only accumulate locally in infected tissues, PR proteins are produced systemically in the plant as part of the host immune response and are associated with pathogen-induced systemic acquired resistance (SAR). As a result, production of PR proteins in uninfected tissue suppresses further pathogen invasion (Van Loon et al., 1987).

PR proteins were first revealed and reported in tobacco mosaic virus (TMV) infected tobacco plants (Van Loon and Van Kammen, 1970). Later, PR proteins were found to be induced in several viroid infected plants such as, tomato and potato. The viroids include several members of the Pospiviroidae family: PSTVd, CEVd, CPFVd, and CSVd (Conejero et al., 1979; Camacho-

Henriquez and Sanger, 1982). To date several host-pathogen interactions have been reported to involve the induction of PR proteins by fungi, bacteria, viruses, and viroids across a broad spectrum of vascular plants (Van Loon, 1997). PR proteins can also be induced in plants by exposure to abiotic stresses such as drought, heavy metals, and ultraviolet rays (Liu and

Ekarmoddoullah, 2006). In addition, PR proteins are constitutive components of many plant tissues (Van Loon and Van Strien, 1999).

Several types of PR proteins are currently recognized, and have been classified into 17 families based on sequence analysis, cellular function and biological activities (Van Loon et al., 2006).

Ten of the seventeen PR families (PR-1, PR-2, PR-3, PR-4, PR-5, PR-8, PR-11, PR-12, PR-13, and

PR-14) have been reported to have antifungal properties. PR-17 proteins have been described as having proteolytic properties analogous to zinc-metalloproteinases (Van Loon et al., 2006).

Pathogen-induced PR-15 and PR-16 proteins are oxalate oxidases with superoxide dismutase activity (Park et. al., 2004) and are putatively associated with signal transduction pathways regulating the hypersensitive response. Members of the PR-12, PR-13 and PR-14 families act as defensins, thionins and lipid-transfer proteins, respectively (Terras et al., 1992; Epple et al.,

1995; García-Olmedo et al., 1995). PR-10 family members have ribonuclease activity, and are

44 the only PR proteins shown to have antiviral properties (Zhou et al., 2002). Proteins from PR-8 and PR-11 families have endochitinase properties capable of chitin hydrolysis. In addition, some

PR-8 proteins have lysozyme activity against bacterial pathogens (Metraux et al., 1989). PR-9 proteins are lignin-forming with peroxidase activity, associated with the oxidative crosslinking of plant cell wall molecules in order to avert pathogen penetration (Lagrimini et al., 1987).

Protection from herbivorous insects and nematodes has been associated to some members of the PR-6 family, which are proteinase inhibitors (Ryan, 1990). Biological activities for PR-5 proteins include membrane permeabilizers and glucan hydrolases (Osmond et al., 2001; Van

Loon et al., 2006). PR-2 and PR-5 proteins, β-1,3-glucanase and thaumatin-like proteins (TLP), respectively; are capable of hydrolyzing glucan present in fungi cell walls. In addition, PR-5 protein P23 is a viroid-induced protein unregulated in tomato plants infected with CEVd

(Rodrigo et al., 1993). P23 has been shown to be an analog of NP24, which is associated with regulating salt stress. Recently, NP24 has also been shown to induce apoptosis in

Saccharomyces cerevisiae (Higuchi et. al., 2015).

Summary and Project goals.

New understandings of the involvement of PR proteins, sRNAs, transcription factors and the different genetic sequences that recognize them have increased dramatically over the past decade. A similar increase has occurred in our understanding of gene regulatory networks and the biochemical pathways they control that are important in disease resistance. This understanding coupled with new high through-put sequencing applications such as RNAseq,

45 and whole genome sequencing has given plant scientist and breeders a new set of tools that can be used in marker-assisted selection strategies for the next set of discoveries that will lead to the genetic improvement of new hop varieties. Such strategies when used to evaluate both coding and non-coding markers specific to a trait allows breeders to evaluate seedlings for a set of desired traits early in the breeding process (Collard and Mackill, 2008). As a result of early trait detection, unwanted plants can be culled and promising plants maintained for future phenotypic evaluation. Such strategies reduce the cost in time and money to the breeding program by reducing the number of plants needed to be maintained from year to year, and by increasing the odds of selecting plants with the most desirable traits. This ability to gain insight into a plants phenotype via early, high through-put DNA-marker detection and the use of advanced molecular biology techniques, to evaluate putative DNA markers as part of a gene regulatory network associated with a specific trait has greatly helped in the development of high yielding, disease and pest resistant plants.

Goals of this thesis project were to combine current understanding of biochemical pathways and gene regulatory networks with high through-put sequencing applications to identify new

DNA-markers that distinguish HSVd-tolerant plants from HSVd-sensitive plants, and to identify potential genes involved in the molecular mechanisms associated with hop stunt disease. Two high through-put sequencing data sets (RNAseq and whole genome sequencing) were produced and compared for gene identification and changes in gene expression patterns. Changes in gene expression patterns were verified by qRT-PCR. Genes shown to have different expression

46 patterns by RNAseq and qRT-PCR were isolated from the whole genome sequencing data and analyzed for sequence variation. In addition, these genes were analyzed in silico for core promoter and cis regulatory elements positioned within a spatial frame work consistent with known functional genes.

RNAseq data sets generated and compared were: 1) Coding and non-coding RNA transcriptomes of HSVd-infected and HSVd-free hop plants.

2) Coding and non-coding RNA transcriptomes, and sRNA data sets between two cultivars of

Humulus lupulus var. lupulus: ‘Nugget’, a HSVd tolerant variety and ‘Glacier’, a HSVd sensitive variety. Whole genome sequencing data sets were generated and compared from six hop cultivars known to have varying degrees of tolerance and sensitivity to HSVd infection. The six cultivars included ‘Cascade’, ‘Columbus’, ‘Galena’, ‘Glacier’, ‘Nugget’, and ‘Willamette’.

Possible outcomes of the combined whole genome sequencing data and RNAseq data sets above would be the putative identification of genes and gene products, including sRNAs that are directly regulated as a result of HSVd infection. Further analysis of the regulated genes and or gene products may provide data that establishes a link between HSVd infection and the molecular mechanism that are directly responsible for tolerant and/or sensitive symptoms observed in different hop cultivars. Such genetic links could be used as DNA markers by hop breeders for marker-assisted selection of new hop cultivars with increased resistance to hop stunt disease.

Prior to the start of this research project a five-year field study of HSVd infected and uninfected hop cultivars ‘Glacier’, ‘Nugget’, ‘Cascade’, ‘Columbus’, ‘Galena’, and ‘Willamette’ was done to compare differences in alpha and beta acid production, cone yield, and plant growth. Results revealed ‘Nugget’ and ‘Galena’ to be the most tolerant hop cultivars to HSVd infection and

‘Glacier’ to be the most severely affected. Three years into the study HSVd infected ‘Glacier’ plants were removed due to the need to control spread of HSVd within the research block.

Average dry cone yield was reduced by 66% in ‘Glacier’ during the first three years, and the average cone weight reduction in ‘Willamette’, ‘Cascade’, ‘Columbus’, ‘Galena ’ and ‘Nugget’ over five years was 28, 14, 6, 6 and 2% respectively. Alpha acid production per acre was reduced by 73% in ‘Glacier’ in the first three years and by 36% and 17% for ‘Willamette’ and

‘Cascade’ respectively. No significant difference in alpha acid production was observed during the five years for ‘Columbus’, ‘Galena’, and ‘Nugget’. Beta acid production per acre was reduced by 76% in ‘Glacier’ in the first three years, and by 36% and 18% for ‘Willamette’ and

‘Cascade’ respectively. No significant difference in beta acid production was observed during the five years for ‘Columbus’, ‘Galena’, and ‘Nugget’. Based on these findings ‘Nugget and

‘Glacier’ were chosen to represent cultivar genotypes that were HSVd tolerant and HSVd sensitive, respectively. In addition during the course of this research project, a survey for the presence of HSVd, using RNA isolated from leaf samples collected on FTA cards from the three major hop growing regions in Washington State was completed in 2011. Much of the work for these two companion projects was completed by others, including Dr. Ken Eastwell, Dr.

Stephen Kenny, and Madhu Kappagantu.

CHAPTER 2

MATERIALS AND METHODS

All hop cultivars were started from virus-free hop plants cultivated in Clean Plant Center

Northwest (CPCNW) facilities located at the Washington State University Irrigated Agriculture

Research and Extension Center (IAREC), Prosser, WA. Plants were propagated from softwood cuttings with one or two nodes. Cuttings, 25 mm to 45 mm long, were surface sterilized by gently agitating in 100 ml of a 10% v/v bleach solution (10 ml Clorox [6.15% w/v sodium hypochlorite], 90 ml ddH2O) for 2 minutes followed by 3 rinses, 3 minutes each in 100 ml ddH2O. Sterilized cuttings were placed into 25 mm X 100 mm sterilized glass tissue culture tubes fitted with breathable caps and containing 20 ml of sterile rooting media (table 1).

Cuttings were grown at 22o C to 25o C on a 14 hour light, 10 hour dark light cycle for 20 to 30 days.

Plantlets with sufficient root development after 20 to 30 days were transferred to 237 ml

Styrofoam planting cups half filled with sterilized potting media (Sunshine Mix # 4 Aggregate

Plus, Sun Gro Horticulture, Agawam, MA) and sealed in ziploc plastic bags. Plants were grown on a 14 hour light, 10 hour dark light cycle at 22o C to 25 o C. After 30 days bags were opened and plants were grown for another 21 days before being transferred to 1 gallon planting pots filled with potting soil (Sunshine Mix # 4 Aggregate Plus). Plants in 1 gallon pots were propagated on a 14 hour light, 10 hour dark light cycle at 30oC to 32oC pre and post inoculation as described above.

Cloning.

Plasmid ‘F56’ is TA Cloning vector pCR2.1 (Invitrogen, Carlsbad, CA) containing an insert consisting of three head-to-tail repeating units of HSVd. HSVd used in the construction of the plasmid was an isolate recovered from a single hop plant (cultivar ‘Glacier’) known to be infected with HSVd. The plasmid was transformed into TOPO TA cloning One Shot Cells

(Invitrogen) according to the manufactures instructions. Briefly, 2l F56 plasmid (10 ng/l) was mixed with one vial of One Shot Chemically Competent E. coli cells, and incubated on ice for 10 minutes followed by a 30 second incubation at 42o C. Following the addition of 250 ml of Super

Optimal broth with Catabolite repression (SOC) (2% w/v tryptone, 0.5% w/v yeast extract, 10 mM NaCl, 2.5 mM KCl, 10 mM MgCl2, 10 mM MgSO4, and 20 mM glucose), the mixture was incubated at 37o C for 1 hour on a horizontal shaker rotating at 200 RPM. Fifty microliters of the transformation reaction was spread evenly across 100 mm2 agar plates containing 25 ml agar prepared using imMEDIA Kan Agar premeasured packets containing 50 g/ml kanamycin

(Invitrogen). Plates were incubated overnight at 37 o C. Several colonies were tested by polymerase chain reaction (PCR) for the presence of vector containing an HSVd polymeric insert as follows: A colony stab of one colony on the end of a micropipette tip was immersed in 18.75

l double distilled water (ddH2O) mixed with 0.75 l 50 mM MgCl2, 2.5 l 10X PCR buffer

(Invitrogen) (200 mM Tris-HCl, (pH 8.4), 500 mM KCl), 0.5 l 10 mM dNTPs, 0.5 l 20 M forward primer HSVd1 (GCC CCG GGG CTC CTT TCT CAG GTA AG), 0.5 l 20 M reverse primer

HSVd2 (GGC AAC TCT TCT CAG AAT CC), 0.5 l 5U/l Taq DNA Polymerase (Invitrogen). HSVd 1

50 and 2 primers are modified version of primers, VP20-H (CGC CCG GGG CAA CTC TTC TCA GAA

TCC), VP19-C (GCC CCG GGG CTC CTT TCT CAG GTA AG) VP20-H corresponds to HSVd2 and

VP19-C to HSVd1 (Kusano and Shimomura, 1997; Astruc et al., 1996). PCR conditions included an initial melt at 94o C 2 for minutes, followed by 35 cycles of: cDNA melting at 94o C for 15 seconds, primer annealing at 62o C for 2 minutes, primer extension at 68o C for 30 minutes, followed by a final primer extension of 68o C for 30 minutes. PCR products were resolved by capillary gel electrophoresis using a QIAxcel instrument (Qiagen, Valencia, CA) equipped with a

12 lane QIAxcel DNA screening cartridge (Qiagen). Samples were run using method AM420.

AM420 loading and running parameters are intended for DNA concentrations of 10-100 ng/l.

Loading parameters were set for a sample injection time of 10 seconds using a voltage of 10 kV.

Separation parameters were set for a run voltage of 5 kV for 420 seconds. Each capillary lane, including a capillary containing a 15bp to 1000bp reference marker were brought into alignment using a QX alignment marker supplied with each cartridge. PCR products showing a positive result for trimeric HSVd insert are indicative that a specific bacterial colony is composed of bacteria containing plasmids with the same trimeric insert.

Identified colonies were cultured overnight at 37o C in 3 ml Luria broth (LB) (10 g tryptone, 5 g yeast extract, 5 g NaCl, 1 ml 1 N NaOH in 1 liter ddH20) with 50 g/ml kanamycin. Plasmids from overnight cultures were isolated using a QIAprep Spin Miniprep kit (Qiagen) and a QIAcube

(Qiagen) instrument according to the manufactures instructions (plasmid DNA Method).

Two milliliters of bacterial overnight cultures were pelleted at 6,800 x g for 3 minutes in safe-

51 lock microcentrifuge tubes. Pelleted cells, reagents, spin columns (DNeasy mini spin column and

QIAshredder spin columns) and disposable filter tips for plasmid purifications were placed into a QIAcube. Reagents included: resuspension buffer P1 (50 mM Tris-HCl pH 8.0, 10 mM EDTA,

100 μg/ml RNAse A), cell lysis buffer P2 (200 mM NaOH, 1% w/v SDS), N3 (4.2 M guanidine-HCl,

0.9 M potassium acetate pH 4.8 ), binding buffer PB (5 M guanidine-HCl, 30% v/v isopropanol), wash buffer PE (10 mM Tris-HCl pH 7.5, 80% v/v ethanol), and elution buffer EB (10 mM Tris-

HCl, pH 8.5). Fifty microliters of purified plasmid in elution buffer EB was collected. Plasmid concentration and quality was measured using a UV-Visible Spectrophotometer (Genesys 10

Bio, ThermoScientific, Waltham, MA) equipped with a nanocell accessory. Fixed wavelength readings were taken at 230, 260, and 280 nm. Plasmid preparations having a 260/280 ratio of

1.6 or greater and a concentration of 180 ng/l or above were saved for insert verification by

PCR and sequencing. Purified plasmid (1 l of 224 to 241 ng/l) was analyzed as described above for colony stab.

Plasmids showing positive PCR results for trimeric HSVd were sent to ELIM Biopharmaceuticals

(Hayward, CA) for sequencing. Sequencing results were compared to know HSVd sequences submitted to the National Center for Biotechnology Information (NCBI) using the NCBI Basic

Local Alignment Search Tool (BLAST) set for highly similar sequence alignment. Aliquots of bacterial colonies grown in overnight cultures that produced plasmids with 100% identity to

HSVd sequences in NCBI were frozen at -80 o C in sterilized 15% v/v glycerol to LB solution for future use.

Cucumber and hop inoculations.

Plasmids containing trimeric head to tail HSVd repeats were isolated as described above and used for inoculating cotyledons of 12 cucumber plants (Cucumis sativus L. cv. Suyo Long) 10 days after emergence. Trimeric plasmids were diluted in 0.1 M Tris-HCl, pH 7.5, to a concentration of 10 g/ml. Cotyledons were dusted lightly with sterilized carborundum, 30 l of dissolved plasmid or mock inoculum (containing buffer void of plasmid) were lightly rubbed into the cotyledon with a fingertip covered with a nitrile glove. Plants were inspected visually for HSVd symptoms, and leaf tissue harvested for RNA extraction 30 days post inoculation.

Total RNA from collected plant tissue was isolated using (Qiagen) RNeasy plant mini kit (Qiagen) and a QIAcube instrument (Qiagen) according to the manufactures instructions (method, RNA plant). One ml of RLT buffer (Qiagen) containing 10 µl -mercaptoethanol and 0.2% v/v

Antifoam Y-30 Emulsion (Sigma, St. Louis, MO) was added to between 0.065 and 0.075 g of leaf tissue in 2 ml screw cap polypropylene microcentrifuge tubes containing one 6.35 mm and three 3.2 mm diameter stainless steel beads. Tubes were fitted with a screw cap sealed by a silicone o-ring and placed into a FastPrep 24 homogenizer (MP Biomedicals, Santa Ana, CA) for

45 seconds at a speed setting of 5.0 m/s. Homogenized tissue was incubated at 56o C for 3 minutes. Six hundred microliters of homogenized and heated tissue was transferred to 2 ml safe-lock microcentrifuge tubes and centrifuged at 14,000 x g for 2 min. RNA was isolated from the tube with the pelleted homogenate via the RNAeasy plant mini kit (Qiagen) in a QIAcube instrument (Qiagen). Seventy microliters of RNase-free H2O containing purified RNA was

53 collected and assayed by RT-PCR for HSVd. RT-PCR reactions using SuperScript III One-Step RT-

PCR System with Platinum Taq DNA Polymerase (Invitrogen) were performed according to the manufactures instructions. Three microliters of sample RNA are added to a master mix buffer containing 12.5 l 2X buffer (Invitrogen), 6 l RNase-free H2O, 0.25 l 20M HSVd1 forward primer, 0.25 l 20M HSVd2 reverse primer, 2 l 7.5 µg/µl non-acetylated BSA and 1 l enzyme mix containing Super Script III Reverse Transcriptase and Platinum Taq DNA Polymerase

(Invitrogen). RT-PCR conditions included a first strand synthesis at 60o C for 30 minutes, followed by melting at 94o C for 2 minutes and 40 cycles of cDNA melting at 94o C for 15 seconds, primer annealing at 62o C for 1 minute, primer extension at 68o C for 1 minute, followed by a final primer extension of at 68o C for 7 minutes. PCR products were resolved by capillary gel electrophoresis using a QIAxcel instrument equipped with a 12 lane QIAxcel DNA screening cartridge (Qiagen) as described above. PCR products indicating a positive result for

HSVd infection in inoculated cucumber plants were sent to ELIM Biopharmaceuticals for sequencing to verify the presence of HSVd and confirm its sequence was unaltered from that contained in the plasmids containing trimeric head-to-tail HSVd repeats used as an inoculum.

Plasmid stocks confirmed to be infectious on cucumber were linearized with NCO I restriction enzyme (Invitrogen). One microgram of trimeric plasmid was incubated at 37o C for 1 hour in a solution containing: 1 l 10 units/l NCO I (Invitrogen), 2 l 10x buffer K (10 mM dithiothreitol ,

100 mM MgCl2, 1 M KCl, 200 mM Tris-HCl, pH 8.5) (Invitrogen), 2 l 0.1% w/v BSA, add ddH2O to bring reaction volume to 20 l. Reaction was terminated by adding 1 l 0.5 M EDTA, 2 l 3

M NaOAc, and 40 l ethanol. Digested plasmid was cooled at -20o C for 30 minutes then pelleted at 14,000 x g for 20 minutes in a microcentrifuge. The supernatant was removed and the pellet air dried for 5 minutes before resuspension in 12 l ddH2O. Linearized plasmids were examined for digestion by agarose gel electrophoresis using a 1.2% w/v agarose gel (SeaKem LE agarose, Lonza, Rockland, ME) containing 10 l/100 ml GelRed (Biotium, Hayward, CA) prepared with tris-base, acetic acid and EDTA (TAE) buffer (40mM Tris, 20mM acetic acid, and

1mM EDTA, pH 8.0). Five microliters of digested or undigested plasmid was mixed with 10 l ddH2O, and 3 l of 6X loading dye (15% w/v Ficoll-400, 66 mM EDTA, 0.012% w/v SDS, 0.09% w/v bromophenol blue, 19.8 mM Tris-HCl, pH 8.0) (New England BioLabs, Ipswich, MA). One microliter of 1 Kb Plus DNA Ladder (Invitrogen) was used for a reference marker. Samples were resolved using a constant 100 volts for 30 minutes and visualized and photographed using a

SYNGENE GBOX equipped with a Chemi HR 16 camera, UV-transilluminator, and GeneSnap imaging software (version 7.12-d).

Linearized plasmids in ddH2O were used for RNA transcription using a mMESSAGE mMACHINE

T7 Ultra Transcription Kit (Ambion, Austin TX) according to the manufactures instructions. One microgram of linearized plasmid in 12 l ddH2O was added to 2 l T7 Enzyme mix (Ambion), 1 l each of 15 mM ATP, CTP, UTP, and 1 l of 3mM GTP, and 2 l 10X T7 reaction buffer (Ambion).

Contents were mixed and heated for 1 hour at 37o C. Unused nucleotides were removed from each transcript preparation with a NucAway spin column (Ambion) according to the manufacture’s instructions. Columns were hydrated with 650 l RNase-free water at room

55 temperature for 15 minutes; excess fluid was removed by centrifugation at 750 x g for 2 minutes. Twenty microliters of prepared transcript was added to each column, eluant free of unused nucleotides was collected by centrifugation at 750 x g for 2 minutes. Clarified transcripts were treated with TURBO DNase (Ambion) according to the manufactures instructions. One microliter of TURBO DNase (2U/l) was mixed with each 20 l transcript reaction and incubated for 15 minutes at 37o C. One microliter from each transcript reaction was removed prior to DNase treatment for comparisons to post DNase treated samples. In addition 1 l of post DNase treated transcript was removed and treated with RNase T1

(Ambion). One microliter of RNase T1 (1U/l) was mixed with 7 l RNase-free water, 1 l of post DNase treated transcript, and 1 l 10X structure buffer (Ambion). The mixture was incubated at room temperature for 30 minutes, followed by heat inactivation at 65o C for 15 minutes. Pre and Post DNase and RNase treated and untreated transcripts were examined by electrophoresis using a 0.8% w/v agarose gel, prepared with MOPS buffer (200mM 3-(N- morpholino) propanesulfonic acid (MOPS), 50 mm NaOAc, 10mM EDTA, pH 7.0) containing

0.74% v/v formaldehyde. Three microliters of sample was mixed with 6 l of denaturation mix

(13 l 37% formaldehyde, 22 l formamide, 65 l 10X MOPS) and resolved using a constant 110 volts for 30 minutes. In addition, RNA transcripts were verified to contain HSVd by RT-PCR.

Reactions using SuperScript III One-Step RT-PCR System with Platinum Taq DNA Polymerase

(Invitrogen) and 1 l of RNA transcript were performed as described above.

Twenty microliters of RNA transcripts freed of unused nucleotides but not DNase treated, was mixed with 4 l RNase free H2O, 30 l 200 ng/l F56 plasmid, and 6 l 0.5 M Tris-HCl, pH 7.5.

Twelve hop plants, of each cultivar, ‘Glacier’ and ‘Nugget’ were kept in the dark 8 to 10 hours prior to inoculation. Six plants from each cultivar received inoculum containing HSVd and six received mock inoculum, free of HSVd. Four leaves approximating a north, south, east, and west location near the base of the plant but not touching the soil were dusted slightly with a small quantity of sterilized carborundum. Thirty microliters of inoculum were lightly rubbed into the leaf with a fingertip covered in a nitrile glove. Plants were inspected visually for HSVd symptoms and leaf tissue harvested for RNA extraction for a period of 30 to 270 days post inoculation.

Hop Rooting Media

Chemical Name Final concentration mg/L

Ammonium nitrate 825 Potassium nitrate 950

Potassium phosphate monobasic 85

Boric acid 3.1

Potassium iodide 0.415

Sodium molybdate dihydrate 0.125

Cobalt(II) chloride hexahydrate 0.0125

Calcium chloride dihydrate 220

Magnesium sulfate heptahydrate 185

Manganese(II) sulfate monohydrate 8.45 Zinc sulfate heptahydrate 4.3 Copper(II) sulfate pentahydrate 0.0125 Ethylenediaminetetraacetic acid disodium salt dihydrate 18.625 Iron(II) sulfate heptahydrate 13.925 Thiamine hydrochloride 0.5 Pyridoxine hydrochloride 0.5 Nicotinic acid 0.5 PABA 2.5 myo-Inositol 0.25 Folic acid 5 Riboflavin 0.05 Ca- Pantothenate 0.25 Biotin 0.5 IBA 2

Table 1. Hop rooting media final concentration in mg/L, pH 5.5.

RNA extractions and high through-put sequencing.

Single leaves matched for size and age were collected from the same and different plants, each leaf was placed into individual plastic bags for transport. Leaves were collected with a single edge razor; new razors were used for each leaf collection, and nitrile gloves were used and changed between each collection. RNA extractions from collected tissue was done using a modified cetyltrimethylammonium bromide (CTAB) method (A. Untergasser

WWW.untergasser.de/lab, accessed October 8, 2014). Briefly, 0.70 to 0.75 g of leaf tissue from a single leaf was placed into 1.8 ml stainless steel microvials pre-chilled on dry ice, containing one 6.35 mm stainless steel bead, three 3.2 mm stainless steel beads and capped with silicone rubber caps (Biospec, Bartlesville, OK). Vials were placed in a FastPrep 24 homogenizer equipped with a CryoPrep adapter (MP Biomedicals) filled with dry ice and agitated for 45 seconds at a 5.0 m/s speed setting. Tubes containing the homogenized tissue was immediately placed on dry ice. Six hundred microliters of CTAB buffer (CTAB 2% w/v, Polyvinylpyrolidone M.

W. 40,000 (PVP 40) 2% w/v, NaCl 1.4 M, EDTA 20 mM, TRIS 100 mM, pH 8.0 plus 1% v/v - mercaptoethanol) was added to each vial followed by the addition of 600 l chloroform. The preparation was mixed by pipetting up and down several times then transferred to new 2 ml safe-lock microcentrifuge tubes, shaken by hand and centrifuged for 2 minutes at 18,400 x g.

The upper aqueous layer (500 l) was transferred to new 2 ml safe-lock microcentrifuge tubes, containing 600 l chloroform, shaken by hand and centrifuged for 2 minutes at 18,400 x g. The upper aqueous layer (330 l) was transferred to new 2 ml safe-lock microcentrifuge tube containing 450 ml 100% ethanol and mixed by pipetting. The solution was pipetted onto an

RNeasy mini spin column (Qiagen) and centrifuged 30 seconds at 8000 X g. Spin column was placed into a new 2 ml collection tube and 350 l RWT buffer (Qiagen) added followed by a 30 second centrifugation at 8000 X g. Flow-through was discarded and 80 l DNase working solution (10 l RNase free, DNase I 1500 Kunitz units /ml, 70 l RDD buffer (Qiagen)) was added to the top of the column and incubated at room temperature for 15 minutes. Following incubation, 350 μl RWT buffer was added to the top of the RNeasy Mini Spin Column and centrifuged for 30 seconds at 8000 x g. Flow-through was collected and reapplied to the top of the same RNeasy Mini Spin Column and centrifuged for 30 seconds at 8000 x g. Five hundred l

RPE (Qiagen) was added to the RNeasy Mini Spin Column and centrifuged for 30 seconds at

8000 x g, followed by the addition of another 500 l RPE into the RNeasy Mini Spin Column, and centrifuge for 2 minutes at 8000 x g. Spin columns were placed into new 2.0 ml collection tubes and centrifuged for 1 minute at 8000 x g to dry the columns. RNA was immediately eluted into a new RNAse free 1.5 ml collection tube by the addition of 70 l RNase free water to the top of spin column and centrifuge for 1 minute at 8000 x g. RNA quality was measured using a UV-

Visible Spectrophotometer equipped with a nanocell accessory (ThermoScientific). Fixed wavelength readings were taken at 230, 260, and 280 nm. RNA preparations having a 260/280 ratio of 1.8 or greater, 230/260 ratio of 1.8 or greater, a concentration of 180 ng/l or above, and a banding pattern consistent of undegraded plant RNA on 1% non-denaturing agarose gels were stored at -70o C.

RNA was tested for the presence of HSVd by RT-PCR reactions using SuperScript III One-Step RT-

PCR System with Platinum Taq DNA Polymerase (Invitrogen) as described above.

Selected RNA samples were submitted for construction of Illumina sequencing libraries with ribosomal RNA depletion using an Illumina TruSeq Stranded Total RNA Sample Prep with Ribo-

Zero (plant) kit (Illumina, San Diego, CA) (University of Utah, Huntsman Cancer Institute, Core

Center for High-Throughput Genomics Salt Lake City, UT). Sequencing of libraries was done with an Illumina HiSeq 2000 sequencer and a HiSeq 50 Cycle Single-Read Sequencing Cluster Kit v4. All library preparations were checked, before sequencing, for adequate quality by evaluation on an Agilent 2100 Bioanalyzer system equipped with an Agilent Bioanalyzer DNA

1000 chip (Alignment Technologies, Santa Clara, CA). Expected library fragment size was 260 base pairs. Raw sequencing data from each library preparation trimmed of adapter sequences was received in an FASTQ file format.

RNA sequence expression analysis and gene discovery.

Raw sequence FASTQ file format data was imported into CLC Genomics Workbench (version

5.1.5) for sequence and alignment analysis. Each sample was analyzed for transcript expression by alignment to three separate hop reference sequences downloaded from NCBI. Hop data bases included: GenBank assembly [GCA_000831365.1] (132,476 scaffolds total sequence length 2,049,208,583), GenBank Transcript BioProject 270039 (387,929 transcripts) and

GenBank Transcript BioProject PRJNA175602 (174,938 transcripts). Number of reads for each

RNAseq sample analyzed ranged from 19,000,000 to 38,000,000.

Expression analysis differences were calculated using the CLC Genomics, RNAseq platform.

Briefly, mapped reads are normalized using Reads per Kilobase per Million mapped reads

(RPKM). RPKM values are compared between samples and a fold change calculated between chosen samples. Samples that showed a greater than two fold change for a specific transcript

(contig) and had a minimum of twenty-five mapped reads were tabulated and saved.

Tabulated transcripts were aligned to the hop reference genome (GenBank assembly

GCA_000831365.1) using NCBI (MEGABLAST) with the highly similar sequence alignment default parameters (initial word size 11 and expectation value 10). Regions of scaffold sequence from the reference genome that showed good alignment to tabulated transcripts were excised and used for further “somewhat similar” BLAST analysis against the entire NCBI nucleotide collection, excluding human and mouse sequences. Scaffolds and specific regions from each scaffold that showed alignment with a plant gene were downloaded to CLC Genomics

Workbench and used as reference sequences for remapping and expression analyses of sample reads. Briefly, entire scaffolds and specific sequence regions within a chosen scaffold were scanned for differences visually and by RNAseq expression analysis with RPKM normalization.

Visual examination was a simple inspection of the graphical representation of individual reads, represented as multiple short straight lines, aligned to scaffold regions, represented as a single long line. Scaffold regions that had clear differences in the number of short lines (reads) aligned to it, had a greater than two fold change between treatments, and had a minimum of twenty-five mapped reads were tabulated and saved. Specific regions from each scaffold that met these parameters were realigned against the entire NCBI nucleotide collection, excluding

62 human and mouse sequences, using a “highly similar sequences” BLAST analysis. Regions that showed good alignment with a plant gene were then mapped back to the original scaffold in its entirety.

Several scaffold regions from the reference genome were identified as having putative plant gene sequence similarities and showed relatively high fold difference between treatment groups. One scaffold, in particular, contained a region which showed good alignment to a well characterized plant defense gene (TLP) composed of a single exon. Further sequence analysis was performed on this plant defense gene sequence to determine if the identified region was composed of attributes associated with a functional gene, e.g. translational start site, full length coding sequence (CDS), core promoter sequence and other regulatory elements associated with gene regulation and function.

Briefly, the putative TLP CDS was scanned using NCBI open reading frame finder for all possible open reading frames and the associated amino acid sequence for the translation product that corresponded with each open reading frame identified. Translation products similar in size to published protein products of the putative TLP protein were chosen for amino acid sequence alignment against the entire NCBI protein data base using NCBI blastp (protein-protein BLAST).

In addition 7,700 bp of upstream sequence (using the translational start site as + 1) from the reference genome was analyzed for core promoter and regulatory elements associated with functional plant genes using “Database of Plant Cis-acting Regulatory DNA Elements” (PLACE).

Identified elements were aligned back to the scaffold to determine each regulatory elements’ relative position to each other and to the translational and transcriptional start sites.

Changes in gene expression between treatment groups of the above identified TLP gene were verified by quantitative Real-Time Polymerase Chain Reaction (qRT-PCR). Briefly, several primer pairs were chosen from a list of primers generated using NCBI Primer-BLAST for qRT-PCR optimization. The TLP CDS from the isolated scaffold region with a complete CDS and associated functional gene components was used for the input sequence for primer generation.

Optimized qRT-PCR conditions produced a PCR product of 196 bp. QRT-PCR reactions utilized reagents from SuperScript III One-Step RT-PCR System with Platinum Taq DNA Polymerase

(Invitrogen) in addition to EvaGreenTM (Biotium), a green fluorescent nucleic acid dye with excitation and emission spectra, when bound to double stranded DNA, of 500 and 530 nm respectively. Briefly, each reaction tube consisted of: 1 l of sample RNA, adjusted to a concentration of 75 ng/l, 24 l of a master mix buffer containing 12.5 l 2X buffer (Ambion),

6.75 l RNase-free H2O, 0.25 l forward primer TLP18F (GTG CCA AGC CTA TGG GCA G), reverse primer TLP18R (CCT CAAC TGA GCA GGG CAT T) 20 m each, 2 l 7.5 µg/µl BSA, non-acetylated,

1 l of enzyme mix containing Super Script III Reverse Transcriptase and Platinum Taq DNA

Polymerase (proprietary) and 1.25 l 20X EvaGreenTM (Biotium). PCR conditions: First strand synthesis, 58o C for 30 minutes, followed by 94o C for 2 minutes and 40 cycles of [melting 94o C for 15 seconds, annealing at 58o C for 30 seconds, extension at 68o C for 1 minute] and a final extension of 68o C for 5 minutes. All qRT-PCR reactions were performed with a Rotor-Gene

6000 instrument equipped with 6000 series software, version 1.7 (build 3) and set to capture fluorescent readings at 520 nm with a gain setting of 7.

To affirm that Real-time quantification was done using qRT-PCR products of the correct size, products from each sample were resolved by capillary gel electrophoresis using high voltage capillary electrophoresis (QIAxcel, Qiagen) equipped with a 12 lane QIAxcel DNA screening cartridge (Qiagen) as described above. In addition, to verify the authenticity of the PCR products, selected qRT-PCR products were sent to ELIM Biopharmaceuticals (Hayward, CA) for direct DNA sequencing.

All gene expression values generated using qRT-PCR were normalized to expression levels of

Glyceraldehyde-3-Phosphate Dehydrogenase (GAPDH) as a reference (Stajner et. al., 2013;

Natsume et. al., 2015). Optimized qRT-PCR conditions for GAPDH produced a PCR product of

119 bp. QRT-PCR reagents and reaction conditions were similar to those described above with the following exceptions: forward primer qRT-HI_GADPH-FW2, (GGG TGG TGC TAA GAA GGT

TG), reverse primer qRT-HI_GADPH-RV2 (TGG TGC AAC TAG CAT TGG AA) and PCR conditions: first strand synthesis, 55o C for 30 minutes, followed by 94o C for 2 minutes and 40 cycles of melting at 94o C for 15 seconds, annealing at 52o C for 30 seconds, extension at 68o C for 1 minute and a final extension of 68o C for 5 minutes. All qRT-PCR reactions were performed with a Rotor-Gene 6000 instrument equipped with 6000 series software, version 1.7 (build 3) and set to capture fluorescent readings at 520 nm with a gain setting of 7. In addition, the TLP CDS region amplified by qRT-PCR was used to search the 132,476 hop reference genome scaffolds

(GenBank assembly GCA_000831365.1) using NCBI (BLASTn) set for somewhat similar sequences alignment default parameters (initial word size 7 and expectation

65 value 10). Scaffold regions that showed alignments to the CDS indicated the possibility that the amplified qRT-PCR products were being generated from other genes, e.g., paralogs or heterozygotic alleles. Each reference scaffold region identified as having sequence similarity was further analyzed to determine if a qRT-PCR product of the correct size and sequence could be generated from identified regions. TLP18F forward primer and TLP18R reverse primer, used in the above TLP qRT-PCR were aligned to each scaffold region using NCBI (BLAST) set for highly similar sequences (initial word size 11 and expectation value 10). Scaffold regions which showed alignments to both primers were mapped to determine relative positions of the primer binding sites, sequence similarities to each primer, and size of PCR product that could be generated.

Sequence variations between hop cultivars.

To determine genetic variation between hop cultivars; ‘Glacier’, ‘Nugget’, ‘Cascade’,

‘Columbus’, ‘Galena’, and ‘Willamette’ DNA was extracted from leaf tissue taken from each cultivar using a DNeasy Plant Mini Kit (Qiagen) according to the manufactures instructions.

Briefly, 0.6 grams of leaf tissue was ground in a 4 X 5 inch mesh grinding bag (Agdia, Elkhart, IN) containing 3 ml AP1 buffer (10 mM Tris-HCl, pH 8.0, 1 mM EDTA pH 8.0, 0.1% SDS, 0.1M NaCl)

(Qiagen), plus 90 l -mercaptoethanol, 30 l RNAse A (100 mg/ml). Six hundred microliters of lysate was transferred to a 2 ml Eppendorf tube and incubated at 65-70o C for 10 minutes. One hundred thirty microliters of AP2 buffer (3.0 M potassium acetate, pH 5.5) (Qiagen) was added to the heated solution and incubated on ice for 5 minutes. The cooled solution was centrifuged at 21,130 X g for 30 seconds. Centrifuged samples were placed into a QIAcube (Qiagen)

66 instrument, for automated DNA extraction using method DNeasy Plant Mini with two 50 l elution steps. Automated pipetting is used in all operations. Briefly, 500 µl of centrifuged sample lysate from individual sample tubes was transferred into separate QIAshredder spin columns (Qiagen) previously placed in the QIAcube rotor adaptor. QIAshredder spin columns are centrifuged for 2 minutes at 6000 X g. Flow-through is transferred to a new tube and 750 µl of buffer AW1(Qiagen) is added and mixed. Six hundred and fifty µl of mixed solution is transferred into a DNeasy Mini spin column (Qiagen) and centrifuge for 1 minute at 6000 X g.

Flow-through is discarded and the step is repeated with remaining sample. Following centrifugation 500 µl of buffer AW2 (Qiagen) is added to the top of the spin column and centrifuged for 1 minute at 6000 X g, flow-through is discarded and the step is repeated with an additional 500µl of buffer AW2 (Qiagen) and a 2 minute centrifugation at 6000 x g . Following washing with buffer AW2 (Qiagen), the DNeasy Mini spin column is transferred and placed within a clean collection tube. Fifty µl of elution buffer AE (Qiagen) is added to the top of the spin column and incubated at room temperature for 5 minutes, followed by a 1 minute centrifugation at 6000 X g. This step was repeated with an additional 50 µl of elution buffer AE

(Qiagen). DNA concentration of eluent and quality was measured using a Genesys 10 Bio UV- visible spectrophotometer equipped with a nanocell accessory (ThermoScientific). Absorbance values were determined at 230, 260, and 280 nm. In addition, each sample was examined by electrophoresis using a 0.8% w/v agarose gel. Three hundred nanograms of DNA in 10 l of

RNAse free water was mixed and added to 2 l of 6X loading dye (0.25% (W/V) bromophenol blue, 0.25% (W/V) xylene cyanol, 40% (W/V) and resolved using a constant 150 volts for 75 to

90 minutes. Two hundred nanograms of DNA from each cultivar in the treatment groups

‘Glacier’ and ’Nugget’, plus DNA from 4 other cultivars (‘Cascade’, ‘Columbus’, ‘Galena’, and

‘Willamette’) were used to the for construction of Illumina sequencing libraries (University of

Utah, Huntsman Cancer Institute, Core Center). Library quality and concentrations are validated on an Agilent 2200 TapeStation. Molarities of libraries are normalized using a qPCR assay (Kapa

Biosystems Library Quantification Kit for Illumina). After normalizing library concentrations, libraries are pooled in preparation for whole genome sequencing using an Illumina HiSeq instrument. Raw sequence FASTQ data was imported into CLC Genomics Workbench (version

5.1.5) for sequence and alignment analysis. Each sample was analyzed by alignment to the hop reference sequence GenBank assembly [GCA_000831365.1] (132,476 scaffolds total sequence length 2,049,208,583). Number of reads for each sample analyzed ranged from 68 to 93 million.

Relative HSVd concentration and TLP expression levels.

Concentration levels of HSVd in infected tissue relative to a control tissue sample were normalized to GAPDH using qRT-PCR and correlated to TLP expression levels.

Optimized qRT-PCR conditions for HSVd produced a PCR product of 180 bp, and utilized reagents from the SuperScript III One-Step RT-PCR System with Platinum Taq DNA Polymerase kit (Invitrogen) in addition to EvaGreenTM (Biotium) as previously described. Briefly, each reaction tube consisted of: 1 l of sample RNA adjusted to a concentration of 75 ng/l, 24 l of a master mix buffer containing 12.5 l 2X buffer (Invitrogen), 8.25 l RNase-free H2O, 0.5 l forward primer HSVd 100 (TTT ACC TTC TCC TGG CTC TTC), reverse primer

HSVdc257 (TTT TCT TTG CTT GCC TTT TG) 20 m each, 1 l of Super Script III Reverse

Transcriptase and Platinum Taq DNA Polymerase (Invitrogen) and 1.25 l 20X EvaGreenTM

(Biotium). PCR conditions were: first strand synthesis 60o C for 30 minute: melting at 94o C for 2 minutes: 40 cycles of melting at 94o C for 15 seconds, annealing at 53o C for 60 seconds, extension at 69o C for 30 seconds and a final extension of 68o C for 5 minutes. All qRT-PCR reactions were performed with a Rotor-Gene 6000 instrument (Corbett Research, Mortlake,

Australia) equipped with 6000 series software, version 1.7 and set to capture fluorescent readings at 520 nm with a gain setting of 7. To affirm that Real-time quantification was done using qRT-PCR products of the correct size each sample was resolved by capillary gel electrophoresis using a QIAxcel instrument (Qiagen) equipped with a 12 lane QIAxcel DNA screening cartridge (Qiagen) as described above.

Survey for the presence of HSVd.

The 2011 National Hop Report stated that ‘Zeus’, ‘Columbus’/’Tomahawk’, ‘Cascade’, ‘Super

Galena’, ‘Willamette’, ‘Apollo’, ‘Nugget’, ‘Bravo’, ‘Chinook’, and ‘Centennial’ were the ten most propagated hop cultivars in Washington State (NASS, 2012). As such these cultivars along with two proprietary cultivars ‘Palisade’, and ‘Simcoe’, and some less commonly propagated cultivars ‘Glacier’, ‘Galena’, ‘Sterling’, and ‘Vanguard’, were surveyed for the presence of HSVd.

Prior to the start of the survey, it was determined that a minimum of 180 samples was needed to reach a 95% confidence interval (Hancock et al., 1993). However, accessibility to each

69 cultivar highly influenced the number of samples collected. As a result of limited accessibility, the number of samples collected varied from 180 for each of the cultivars tested.

During the last week in June and first week in July, 2012, a total of 1635 samples from 20 different hop cultivars were collected from the three major hop growing regions in Washington

State: lower Yakima valley, Moxee Valley and Toppenish creek. Sample collection consisted of collecting and pressing four leaves from each plant onto Whatman™ FTA™ Plant Saver Cards

(Fisher Scientific, Pittsburg PA). Following collection, FTA cards were stored at -20o C until used for RNA extraction.

RNA extraction from FTA cards and RT-PCR for the presence of HSVd (performed by Madhu

Kappagantu).

Four or five squares approximately 2 mm per side were excised from FTA cards with a single edged razor. A new razor was used for each FTA card. Excised squares were placed into 1.5 ml plastic centrifuge tubes and 250 µl of GEB buffer (0.01M Na2CO3, 0.03 M NaHCO3, 2% PVP, 0.5%

Tween-20, 0.2% BSA) was added. Tubes containing FTA squares and GEB buffer were incubated at room temperature for 60 minutes. Following incubation 8 µl of solution was transferred into clean 1.5 ml centrifuge tubes containing 100 µl of GES buffer (0.1 M glycine, pH 8 50mM NaCl,

1mM EDTA, 0.5% Trition X 100) with 1 % v/v -mercaptoethanol. The mixture was incubated at

95oC for 10 minutes followed by rapid cooling on ice. Samples were tested for presence of HSVd by RT-PCR as described above.

CHAPTER 3

RESULTS AND DISCUSSION

Infectious clone of HSVd.

PCR of plasmids isolated from bacterial colonies transformed with plasmid ‘F56’ yielded amplification products consistent in size with HSVd (figure 1). Additionally, sequencing results from purified plasmids showed a single genotype of three head to tail repeating units of HSVd within the plasmid cloning site (figure 2).

Figure 1. Capillary gel electrophoresis of PCR amplification products using plasmid F56 purified from transformed One Shot Cells as a template. Lane 1, ladder (15 – 1000 bp); Lanes 2-4, F56 purified plasmids; Lane 5, HSVd (+) control; Lane 6, negative control (empty vector pCR2.1). Arrow indicates correct amplicon size (297bp).

TGCAGAATTCGCCCTTGCCCCGGGGCTCCTTTCTCAGGTAAGTACCTTCCTGCCT TGTTTTTTTCTTTGCTTGCCTTTTGCGGCAACTCGAGAATTCCCCAGAGGGGCTC AAGAGAGGTTCCGCGGCAGAGGCTCAGATAGACAAAAAGCAGGTTGGGACGAACC GAGAGGTGATGCCACCGGTCGCGTCTCATCGGAAGAGCCAGGAGAAGGTAAAGAA GAAGGGACGATCGATGGTGTTTCGAAGGCAGAGCCTCTACTCCAGAGCACCGCGG CCCTCTCTCCACGCCTCTCGCTGGATTCTGAGAAGAGTTGCCCCGGGGCTCCTTT CTCAGGTAAGTACCTTCCTGCCTTGTTTTTTCTTTGCTTGCCTTTTGCGGCAACT CGAGAATTCCCCAGAGGGGCTCAAGAGAGGTTCCGCGGCAGAGGCTCAGATAGAC

AAAAAGCAGGTTGGGACGAACCGAGAGGTGATGCCACCGGTCGCGTCTCATCGGA

AGAGCCAGGAGAAGGTAAAGAAGAAGGGACGATCGATGGTGTTTCGAAGGCAGAG

CCTCTACTCCAGAGCACCGCGGCCCTCTCTCCACGCCTCTCGCTGGATTCTGAGA AGAGTTGCCCCGGGGCTCCTTTCTCAGGTAAGTACCTTCCTGCCTTGTTTTTTCT TTGCTTGCCTTTTGCGGCAACTCGAGAATTCCCCAGAGGGGCTCAAGAGAGGTTC CGCGGCAGAGGCTCAGATAGACAAAAAGCAGGTTGGGACGAACCGAGAGGTGATG CCACCGGTCGCGTCTCATCGGAAGAGCCAGGAGAAGGTAAAGAAGAAGGGACGAT CGATGGTGTTTCGAAAGGCAGAGCCTCTACTCCAGAGCACCGCGGCCCTCTCTCC ACGCCTCTCGCTGGATTCTGAGAAGAGTTGCCAGGGCGAATTCCAGC

Figure 2. Sequence of F56 plasmid. Boxes outline borders of pCR2.1 plasmid insert site. Alternating bold and regular font represents HSVd monomers.

Sequencing results were compared to know HSVd sequences using NCBI (BLAST). Results indicate that HSVd monomers in F56 are identical to HSVd isolate hKFKi, a well characterized

HSVd isolated from hop (figure 3) (GenBank GI:12082110).

hKFKi 1 CTGGGGAATTCTCGAGTTGCCGCAAAAGGCAAGCAAAGAAAAAACAAGGCAGGAAGGTAC 60 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| F56 1 CTGGGGAATTCTCGAGTTGCCGCAAAAGGCAAGCAAAGAAAAAACAAGGCAGGAAGGTAC 60

hKFKi 61 TTACCTGAGAAAGGAGCCCCGGGGCAACTCTTCTCAGAATCCAGCGAGAGGCGTGGAGAG 120 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| F56 61 TTACCTGAGAAAGGAGCCCCGGGGCAACTCTTCTCAGAATCCAGCGAGAGGCGTGGAGAG 120

hKFKi 121 AGGGCCGCGGTGCTCTGGAGTAGAGGCTCTGCCTTCGAAACACCATCGATCGTCCCTTCT 180 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| F56 121 AGGGCCGCGGTGCTCTGGAGTAGAGGCTCTGCCTTCGAAACACCATCGATCGTCCCTTCT 180

hKFKi 181 TCTTTACCTTCTCCTGGCTCTTCCGATGAGACGCGACCGGTGGCATCACCTCTCGGTTCG 240 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| F56 181 TCTTTACCTTCTCCTGGCTCTTCCGATGAGACGCGACCGGTGGCATCACCTCTCGGTTCG 240

hKFKi 241 TCCCAACCTGCTTTTTGTCTATCTGAGCCTCTGCCGCGGAACCTCTCTTGAGCCCCT 297 ||||||||||||||||||||||||||||||||||||||||||||||||||||||||| F56 241 TCCCAACCTGCTTTTTGTCTATCTGAGCCTCTGCCGCGGAACCTCTCTTGAGCCCCT 297

Figure 3. Sequence of monomer from F56 plasmid aligned to hKFKi a well characterized HSVd isolated from hop.

Plasmid F56 was used for inoculating cotyledons of cucumber plants (Cucumis sativus L. cv.

Suyo Long) 10 days after emergence. Plants started to exhibit symptoms (stunting and chlorosis) 10 to 15 days post inoculation. Mock inoculated plants were free of symptoms (figure

4).

Figure 4. (Left) Mock inoculated cucumber (Cucumis sativus L. cv. Suyo Long). (Right) Cucumber (Cucumis sativus L. cv. Suyo Long) inoculated with plasmid F56 containing a trimeric clone of Hop stunt viroid 45 days post inoculation.

RT-PCR of total RNA collected from fresh cumber leaf tissue produced amplification products of the correct size (figure 5).

Figure 5. Capillary gel electrophoresis of RT-PCR amplification products using total RNA

extracted from cucumber leaf tissue. Lane 1, ladder (15 – 1000 bp); Lanes 2 - 4 inoculated cucumber; Lane 5 mock inoculated cucumber; Lane 6, HSVd (+) control. Arrow indicates correct amplicon size (297bp).

PCR products indicating a positive result for HSVd infection from inoculated cucumber plants were sent to ELIM Biopharmaceuticals for sequencing to verify the sequence was unaltered from that contained in the plasmids used as an inoculum. Results indicate that HSVd sequences amplified from inoculated cucumber plants was identical to hKFKi (figure 6.)

PCR Prd 1 CGCGGTGCTCTGGAGTAGAGGCTCTGCCTTCGAAACACCATCGATCGTCCCTTCTTCTTT 60 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| hKFKi 126 CGCGGTGCTCTGGAGTAGAGGCTCTGCCTTCGAAACACCATCGATCGTCCCTTCTTCTTT 185

PCR Prd 61 ACCTTCTCCTGGCTCTTCCGATGAGACGCGACCGGTGGCATCACCTCTCGGTTCGTCCCA 120 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| hKFKi 186 ACCTTCTCCTGGCTCTTCCGATGAGACGCGACCGGTGGCATCACCTCTCGGTTCGTCCCA 245

PCR Prd 121 ACCTGCTTTTTGTCTATCTGAGCCTCTGCCGCGGAACCTCTCTTGAGCCCCT 172 |||||||||||||||||||||||||||||||||||||||||||||||||||| hKFKi 246 ACCTGCTTTTTGTCTATCTGAGCCTCTGCCGCGGAACCTCTCTTGAGCCCCT 297

PCR Prd 173 CTGGGGAATTCTCGAGTTGCCGCAAAAGGCAAGCAAAGAAAAAACAAGGCAGGAAGGAA 232 ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| hKFKi 1 CTGGGGAATTCTCGAGTTGCCGCAAAAGGCAAGCAAAGAAAAAACAAGGCAGGAAGGTA 59

PCR Prd 233 CTTACCTGAGAAAGGAGCCCCGGGGCAACTCTTCTCAGAATCCAGCGAGAGGCGTGGAGA 291 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| hKFKi 60 CTTACCTGAGAAAGGAGCCCCGGGGCAACTCTTCTCAGAATCCAGCGAGAGGCGTGGAGA 119

PCR Prd 292 GAGGGCCGCGGTGCTCTGGAGTAGAGGCTCTGCCTT 327 |||||||||||||||||||||||||||||||||||| hKFKi 120 GAGGGCCGCGGTGCTCTGGAGTAGAGGCTCTGCCTT 155

Figure 6. Sequence alignment of HSVd RT-PCR product produced using total RNA extracted from F56 inoculated cucumber as a template and HSVd isolate hKFKi.

Tissue culture derived plants of ‘Glacier’ and ‘Nugget’ were inoculated with crude extracts of infected cucumber leaves. HSVd inoculated ‘Glacier’ plants started showing HSVd symptoms 60 to 140 days post inoculation. ‘Nugget’ plants were void of HSVd symptoms for the entire one year of observation (figure 7, 8, and 9).

All plants were tested by PCR for the presence of Hop mosaic virus, Hop latent virus, American hop latent virus, Apple mosaic virus, and Hop latent viroid (HLVd). All plants tested negative for each of the viruses list. All plants tested positive for the presence of HLVd. However, HLVd produces no known symptoms on varieties grown in the Pacific Northwest.

1 2

Figure 7. ‘Glacier’ plants 140 days post inoculation. Plant 1 mock inoculated. Plant 2 HSVd inoculated.

1 2

Figure 8. ‘Nugget’ plants 140 days post inoculation. Plant 1 mock inoculated. Plant 2 HSVd inoculated.

1 2 3 4

Figure 9. ‘Nugget’ and ‘Glacier’ plants 140 days post inoculation. Plants 1 and 2 HSVd inoculated ‘Nugget’. Plant 3 and 4 HSVd inoculated ‘Glacier’.

High Through-Put Sequencing

Single leaves matched for size and age were collected for total RNA extractions from HSVd inoculated and mock inoculated plants between 30 and 270 days post inoculation (figure 10).

To check for quality, each RNA extraction was resolved on a 1% non-denaturing agarose gel

(figure 11). Only RNA preparations having a banding pattern consistent of undegraded plant

RNA and a concentration of 180 ng/l or above, as measured with a UV Spectrophotometer, were saved for further analysis.

Figure 10. Leaves 1 and 2, mock inoculated ‘Nugget’. Leaves 3 and 4, HSVd inoculated ‘Nugget’.

marker

10000

6000 1 2 3 4 4000 3000

2000

1500 1000

500

200 bp

Figure 11. RNA extraction resolved on a 1% non-denaturing agarose gel. Lanes 1 and 2, RNA from HSVd inoculated ‘Nugget’; 349 and 255 ng/µl respectively. Lanes 3 and 4,

RNA from mock inoculated ‘Nugget’; 387 and 224 ng/µl respectively.

Selected RNA samples sent for high-throughput sequencing were ribosomal RNA depleted before construction of Illumina sequencing libraries and each library preparation was checked, for adequate quality by evaluation on an Agilent 2100 Bioanalyzer. Only libraries having the

81 expected library fragment size of approximately 260 base pairs were sequenced. Raw sequence data containing between 19,000,000 to 40,000,000 50 bp reads for each sample were analyzed for transcript expression by alignment to three separate hop reference sequences. Results showed that several genes from the ‘Glacier’ and ‘Nugget’ treatment groups were differentially expressed relative to mock inoculated controls (table 1). A treatment group consisted of HSVd inoculated and mock inoculated plants of the same cultivar.

Cultivar Change ID Sequence alingment BLAST searh result NCBI accession Nugget 2 331881 Brassica napus thaumatin-like protein 1 (LOC106440536), mRNA XM_013882235 Glacier -12.90 Nugget -2.1 332397 Populus euphratica 60S ribosomal protein L18-2 XM_011038594.1 Glacier 2.2 Nugget 2.7 343934 Humulus lupulus vps gene for valerophenone synthase AB047593.2 Glacier 3.3 Nugget 2.4 346875 Jatropha curcas transcription factor bHLH130-like transcript variant X2 XM_012210134.1 Glacier 3.0 Nugget 3.9 350263 Morus notabilis (RS)-norcoclaurine 6-O-methyltransferase XM_010095995.1 Glacier -2.9 Nugget -3.8 352748 Morus notabilis Acyl carrier protein 1 XM_010101745.1 Glacier 2.4 Nugget -5.0 353439 Morus notabilis Preprotein translocase subunit SECE1 XM_010091374.1 Glacier -4.0 Nugget -3.3 355439 Morus notabilis Peroxiredoxin Q XM_010091374.1 Glacier 2.2 Nugget 11.1 357517 Morus notabilis Inositol-3-phosphate synthase XM_010092256.1 Glacier -2.4 Nugget 2.1 358677 Vitis vinifera uncharacterized LOC100254241 XM_002277774.3 Glacier 2.4 Nugget 2.7 359077 Morus notabilis Protein brittle-1 XM_010094931.1 Glacier 3.0 Nugget 3.6 359990 Morus alba var. multicaulis voucher Yu 71-1 CRT/DRE binding factor 1 JX186750.1 Glacier 5.2 Nugget -9.0 363597 Morus alba var. multicaulis voucher Yu 71-1 CRT/DRE binding factor 1 JX186750.1 Glacier 2.3 Nugget -2.8 365362 Morus notabilis S-receptor-like serine/threonine-protein kinase XM_010107312.1 Glacier 2.6 Nugget -2.5 366502 Prunus mume photosystem II repair protein PSB27-H1, chloroplastic XM_008232036.1 Glacier 3.0 Nugget 3.7 374279 Citrus sinensis putative peptidyl-tRNA hydrolase PTRHD1-like XM_006483490.1 Glacier 2.8 Nugget 2.7 376110 Morus notabilis Disease resistance RPP13-like protein 4 XM_010115099.1 Glacier 2.2 Nugget 2.0 380114 Populus euphratica transcription factor bHLH130-like XM_011009047.1 Glacier 3.9 Nugget -4.0 Table 1. Relative387765 fold changePentatricopeptide between repeat -BLASTcontaining identified protein At5g50390, sequences chloroplastic in HSVd infectedXM_008232554.1 and mock inoculatedGlacier -plants2.8 from ‘Nugget’ and ‘Glacier’ treatment groups. Differences were calculated Nugget 3.0 392090 Prunus mume pentatricopeptide repeat-containing protein At2g01390 XM_008243399.1 usingGlacier reads 3.0per kilobase of transcript per million mapped reads (RPKM) values. Negative numbers Nugget 3.1 393521 Prunus mume scarecrow-like protein 21 XM_008246105.1 indicateGlacier a gene2.9 down regulation.

Cultivar Change ID Sequence alingment BLAST searh result NCBI accession Nugget 7.0 393833 Fragaria vesca J domain-containing chloroplast accumulation response 1 XM_004293172.2 Glacier 2.2 Nugget -6.0 397378 Morus notabilis Thioredoxin-like 1-2 XM_010095297.1 Glacier 3.0 Nugget 3.0 397581 Vitis vinifera E3 ubiquitin-protein ligase Arkadia XM_002283506.3 Glacier -2.5 Nugget 7.0 401809 Prunus mume protein Dr1 homolog XM_008245840.1 Glacier 2.5 Nugget -2.4 402568 Morus notabilis Annexin D4 XM_010097646.1 Glacier 2.7 Nugget 5.6 403240 Vitis vinifera tetraspanin-2 XM_002282361.3 Glacier 3.5 Nugget 2.5 405069 Prunus mume probable steroid-binding protein 3 XM_008222425.1 Glacier -2.7 Nugget 2.8 409491 Morus notabilis Zinc finger CCCH domain-containing protein 29 XM_010103125.1 Glacier 6.0 Nugget 3.5 410160 Eucalyptus grandis ubiquitin-like protein 5 XM_010033688.1 Glacier 3.0 Nugget 2.6 414990 Vitis vinifera protein TIFY 10A XM_002272327.3 Glacier 7.2 Nugget -5.3 415383 Fragaria vesca subsp. vesca methylesterase 17-like XM_004294671.2 Glacier 2.5 Nugget -2.7 415386 Fragaria vesca subsp. vesca methylesterase 17-like XM_004294671.2 Glacier 2.3 Nugget 2.0 416722 Rosa chinensis putative TCP9 protein mRNA, complete cds KP784450.1 Glacier 2.7 Nugget -3.3 417808 Fragaria vesca subsp. vesca DEAD-box ATP-dependent RNA helicase 42 XM_004291453.2 Glacier -2.3 Nugget -2.1 424294 Morus notabilis Zinc finger protein XM_010101442.1 Glacier 2.2 Nugget 3.4 426767 Fragaria vesca subsp. vesca transcription factor bHLH18-like XM_011468190.1 Glacier 2.5 Nugget 3.6 427165 Morus notabilis putative E3 ubiquitin-protein ligase RHA2B XM_010109050.1 Glacier 2.6 Nugget 2.4 427347 Morus notabilis Zinc/RING finger protein 3 XM_010100775.1 Glacier 2.9 Nugget 3.6 428584 Prunus mume transcription factor bHLH36-like XM_008230776.1 Glacier 13.5 Nugget 2.6 433716 Fragaria vesca ethylene-responsive transcription factor ERF109-like XM_004294761.2 Glacier 3.3 Nugget 2.4 440001 Prunus mume probable galactinol--sucrose galactosyltransferase 5 XM_008232336.1 Glacier 3.4 Nugget 7.0 443959 Morus alba var. multicaulis transport inhibitor response 1 (TIR1) KJ787017.1 Glacier -2.4 Nugget 5.9 444112 Morus notabilis Benzyl alcohol O-benzoyltransferase XM_010112087.1 Glacier 2.4 Nugget 3.3 446402 Vitis vinifera sarcoplasmic reticulum histidine-rich calcium-binding protein XM_003631163.2 Glacier 4.0 Nugget -2.4 451858 Prunus mume pentatricopeptide repeat-containing protein At5g39980 XM_008232489.1 Glacier 2.6 Nugget -5.0 457909 Morus notabilis DEAD-box ATP-dependent RNA helicase 21 XM_010094901.1 Glacier 2.2 Nugget -6.0 460523 Morus notabilis ABC transporter G family member 26 XM_010110964.1 Glacier -2.6

Table 1. Continued.

Cultivar Change ID Sequence alingment BLAST searh result NCBI accession Nugget 3.4 481219 Fragaria vesca subsp. vesca beta-amylase 1, chloroplastic (LOC101312918), mRNAXM_004296501.2 Glacier 3.1 Nugget 5.0 481847 Prunus mume E3 ubiquitin-protein ligase RNF185-like XM_008243396.1 Glacier -3.0 Nugget 3.5 487103 Morus notabilis Ras-related protein RABA1f XM_010112845.1 Glacier 3.5 Nugget 8.0 490764 Morus notabilis Rho GDP-dissociation inhibitor 1 XM_010114519.1 Glacier 3.5 Nugget 2.7 493776 Morus notabilis NADP-dependent alkenal double bond reductase P2 XM_010103045.1 Glacier 13.4 Nugget 3.8 503582 Morus notabilis Ferric reduction oxidase 4 XM_010107453.1 Glacier -2.2 Nugget 4.0 524444 Fragaria vesca subsp. vesca ras-related protein RABF2a XM_004293684.2 Glacier 4.0 Nugget 3.2 530502 Malus x domestica transcription factor MYB59 XM_008359549.1 Glacier 3.2 Nugget 2.1 530804 Prunus mume sufE-like protein 2 XM_008223670.1 Glacier 2.5 Nugget 4.4 539907 Prunus mume myosin-9-like XM_008242142.1 Glacier 2.9 Nugget 4.0 554653 Theobroma cacao Alpha/beta-Hydrolases superfamily XM_007027657.1 Glacier 3.0 Nugget 2.3 577200 Morus notabilis Zeatin O-xylosyltransferase XM_010091895.1 Glacier 2.7 Nugget -2.4 585022 Morus notabilis putative pectinesterase/pectinesterase inhibitor 51 XM_010092564.1 Glacier 3.4 Nugget 2.6 589572 Morus notabilis Leucine-rich repeat receptor-like tyrosine-protein kinase XM_010096566.1 Glacier 3.7 Nugget 4.6 592583 Morus notabilis Multiple C2 and transmembrane domain-containing protein 1 XM_010107658.1 Glacier 4.5 Nugget 4.2 600524 Populus euphratica patellin-3 XM_011033203.1 Glacier 4.0 Nugget -5.2 601745 Morus notabilis Monothiol glutaredoxin-S2 XM_010100030.1 Glacier 2.1 Nugget 3.1 609857 Morus notabilis Receptor-like protein kinase HAIKU2 XM_010106243.1 Glacier 2.6 Nugget 3.8 610724 Malus x domestica beta-1,4-mannosyl-glycoprotein 4-beta-N-acetylglucosaminyltransferaseXM_008341874.1 Glacier 3.3 Nugget 2.9 617996 Morus notabilis Putative U-box domain-containing protein 42 XM_010092961.1 Glacier 2.2 Nugget -3.9 620372 Fragaria vesca subsp. vesca fatty-acid-binding protein 1 XM_004292542.2 Glacier 2.6 Nugget 3.0 641610 Prunus mume probable nucleoredoxin 2 XM_008222990.1 Glacier 4.0 Nugget -3.0 644601 Morus notabilis putative mitochondrial chaperone bcs1 XM_010115100.1 Glacier 2.1 Nugget -5.0 645548 Morus notabilis Carbonic anhydrase XM_010110474.1 Glacier 2.3 Nugget 2.3 670157 Prunus mume scarecrow-like protein 3 XM_008229016.1 Glacier -3.0 Nugget -2.5 683615 Morus notabilis 30S ribosomal protein S13 XM_010093037.1 Glacier 2.1 Nugget 6.0 685103 Morus notabilis tRNA wybutosine-synthesizing protein 1-like protein XM_010098081.1 Glacier 2.4

Table 1. Continued.

TLP discovery and expression differences between treatment groups.

NCBI BLAST search using the 329 bp contig (LA331881) listed in table 1 above showed a partial alignment of 58 bp (269 to 327) to Brassica napus thaumatin-like protein (TLP) NCBI accession number XM_013882235 (figure 12).

LA331881 TAGTTAGTGCCTCCAGGGCACGT-AAATGTGCTGGTGGGATCATCCTTAGGGTAACTATA A) ||||| || || | |||||| || ||| | |||||||||||||||||||||||||||||| TLP B.napus TAGTTGGTTCCGCTAGGGCAAGTGAAAAG-GCTGGTGGGATCATCCTTAGGGTAACTATA

LA331881 TAGTTAGTGCCTCCAGGGCACGTAAATGTGCTGGTGGGATCATCCTTAGGGTAACTATA B) ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| LD139544 TAGTTAGTGCCTCCAGGGCACGTAAATGTGCTGGTGGGATCATCCTTAGGGTAACTATA

Figure 12. A) Alignment of 58 bp region of contig LA331881 to Brassica napus thaumatin-like protein. B) Alignment of LA331881 putative TLP region to scaffold LD139544.

Alignment of the 58 bp sequence similar to TLP from contig LA331881 against scaffolds from the hop reference genome GenBank assembly GCA_000831365.1 identified a TLP similar region on scaffold LD139544 (figure 12). High through-put sequencing analysis using RPKM expression values from reads mapped to the TLP identified region of LD139544 from 29612 to 30282 indicated a down regulation of TLP in ‘Glacier’ in response to HSVd whereas expression of TLP in ‘Nugget’ exhibited a slight fold increase in response to infection with HSVd (table 2).

Sample Sample Sample Sample Sample Sample 4 5 6 Mapped 1 2 3 Mapped Mapped Mapped Mapped reads Mapped Mapped Mapped reads reads reads reads HSVd RPKM reads reads reads HSVd RPKM HSVd HSVd HSVd free HSVd TLP fold Cultivar HSVd HSVd HSVd total HSVd free free free total free change

'Glacier' 31 31 20 82 0.38 376 138 269 783 4.65 -12

'Nugget' 20 1.0 15 36 0.23 13 21 5.0 39 0.26 1.1

Table 2. Relative fold change for TLP between, HSVd infected (HSVd) and mock inoculated (HSVd free) plants from ‘Glacier’ and ‘Nugget’ treatment groups. FoFoldld changes calculated

using RPKM expression values from high through-put sequencing data for reads mapped to the TLP identified region of LD139544. ‘Glacier’ shows a down regulation of TLP expression and ‘Nugget’ shows little change in TLP expression.

Further analysis of TLP expression levels by qRT-PCR normalized to GAPDH showed a significant difference using a Student two tailed T test and 95% confidence interval for the ‘Glacier’ treatment group (P value 0.00036). However, the difference (2.6) was lower than that calculated using RPKM expression values. Differences between ‘Nugget’ treatment groups showed no statistical significance (P value 0.48) using a Student two tailed T test and 95% confidence interval (tables 3 and 4).

'Nugget' Normalized Normalized expression values HSVd expression values HSVd 0.02 pos 0.04 neg 0.02 pos 0.05 neg 0.02 pos 0.05 neg 0.02 pos 0.05 neg 0.03 pos 0.06 neg 0.03 pos 0.06 neg 0.04 pos 0.06 neg 0.04 pos 0.06 neg 0.04 pos 0.09 neg 0.04 pos 0.13 neg 0.04 pos 0.15 neg 0.07 pos 0.28 neg 0.10 pos 0.28 neg 0.10 pos 0.39 neg 0.11 pos 0.43 neg 2.66 pos 0.50 neg 0.18 pos 1.32 neg 0.18 pos 1.78 neg 0.19 pos 1.90 neg 0.31 pos 0.01 neg 0.50 pos 0.02 neg

0.12 pos 0.04 neg 0.01 pos 0.04 neg

0.03 neg

AVG 0.21 AVG 0.33 STD 0.55 STD 0.54

P value 0.48

Table 3. Results from qRT-PCR, normalized to GAPDH showed no change between HSVd infected and mock inoculated ‘Nugget’ plants. No statistical significance was determined using a Student two tailed T test and 95 % confidence interval.

'Glacier' Normalized Normalized expression values HSVd expression values HSVd 0.10 pos 0.49 neg 0.10 pos 0.51 neg 0.13 pos 0.68 neg 0.13 pos 0.83 neg 0.13 pos 0.92 neg 0.14 pos 0.92 neg 0.15 pos 1.07 neg 0.16 pos 1.17 neg 0.18 pos 1.66 neg 0.28 pos 0.49 neg 0.30 pos 0.58 neg 0.37 pos 0.58 neg 0.38 pos 0.61 neg 0.39 pos 0.63 neg 0.47 pos 0.71 neg 0.51 pos 0.72 neg

0.65 pos 0.72 neg 0.72 pos 0.75 neg

0.94 pos 0.75 neg 1.01 pos 1.48 neg

1.04 pos 2.0 neg 0.05 pos 3.13 neg

AVG 0.38 AVG 0.97

STD 0.31 STD 0.62

P value 0.00036

Table 4. Results from qRT-PCR, normalized to GAPDH showed a 2.6 fold change (down regulation) for HSVd infected ‘Glacier’ plants compared to mock inoculated plants. Statistical significance was determined using a Student two tailed T test and 95 % confidence interval.

Relative HSVd concentration and TLP expression levels.

Concentration levels of HSVd in infected tissue relative to a control tissue sample were correlated to TLP expression levels, normalized to GAPDH using qRT-PCR. Results indicate that changes in TLP expression levels between infected tissue samples from ‘Glacier’ and ‘Nugget’ are independent of HSVd levels (Table 5). Statistical significance was determined using a

Student two tailed T test and 95% confidence interval (P value 0.53).

Table 5. Concentration levels of HSVd in infected tissue correlated to TLP expression levels, normalized to GAPDH using qRT-PCR. Results indicate that changes in TLP expression levels between infected tissue samples from ‘Glacier’ and ‘Nugget’ are independent of HSVd. Statistical significance was determined using a Student two tailed T test and 95% confidence interval.

Additionally, comparative qRT-PCR results of normalized TLP expression levels to HSVd levels within the HSVd infected subset of plants in each treatment group showed a positive correlation for HSVd infected ‘Nugget’ plants and a negative correlation for HSVd infected

‘Glacier’ plants using simple linear regression analysis. Coefficients of determination respectively are 0.351 and 0.356 (figure 13).

Figure 13. Simple linear regression analysis of qRT-PCR results showing the correlation between normalized TLP levels and HSVd levels relative to an HSVd standard for ‘Glacier’ and ‘Nugget’ HSVd infected plants. A) Negative correlation for HSVd infected ‘Glacier’ plants. B) Positive correlation for HSVd infected ‘Nugget’ plants.

TLP coding sequence and sequence analysis.

The TLP region identified in LD139544 was used as a reference sequence for in silico analysis and discovery of the TLP coding sequence (figure 14). NCBI open reading frame software identified an open reading frame and coding sequence (CDS) that coded for a similar number of nucleotides as other known TLPs (figure 14). PBLAST analysis of the CDS identified it as having a similar number of amino acids specific to the plant TLP subfamily (figure 15).

29612 ATGAGGTCCT CTATTATTTT CTCATTTCTT TTAGTGCTAA CTTACTTCTC CGCTTCAACC 29672 CATGCAGCAA GATTCGACAT CACAAACAGA TGCCCCTTCA CCGTGTGGGC AGCTGCCGTG 29732 CCCGGCGGTG GAAAACAGCT GAGCTCGGGC CAATCATGGG CCCTTGACGT CAACGCAGGC

29792 ACGACAGGGG CTCGCATATG GGCTCGGACG AATTGTAATT TTGATGGGGC TGGACGCGGC 29852 AGGTGTGAGA CTGGCGACTG CGGTGGCGTT CTCCAGTGCC AAGCCTATGG GCAGGCGCCC 29912 AATACCTTGG CCGAGTACGC GCTGAACCAA TTCAATAACT TAGATTTCTT TGACATCTCG 29972 CTGGTGGATG GGTTCAATGT CCCCATGGAC TTCAGTCCCA CTTCACCCCA GTGCAGCCGA 30032 GGGATCAAGT GTGTGGCAAA TATAAATAAC GAATGCCCTG CTCAGTTGAG GGCCCCTGGA

30092 GGCTGCAAGG ATCCATGTAA TGTCTTTAAA ACTGATATGT ATTGTTGTAA CTCAGGTAGC 30152 TGTGGACCCA CAGAATTCTC CAAGTTCTTT AAGCAACGAT GCCCTGATGC TTATAGTTAC 30212 CCTAAGGATG ATCCCACCAG CACATTTACG TGCCCTGGAG GCACTAACTA TAGGGTTGTC 30272 TTCTGCCCTT GA

Figure 14. Identified coding sequence from LD139544 region 29612 to 30283 using NCBI open reading frame software. NCBI BLAST predicted Thaumatin-Like Protein coding sequence.

A) MRSSIIFSFLLVLTYFSASTHAARFDITNRCPFTVWAAAVPGGGKQLSSGQSWALDVNAGTTGGA R I WARTNCNFDGAGRGRCETGDCGGVLQCQAYGQAPNTLA E YALNQFNNL D FFDISLV D GFNVP

MDFSPTSPQCSRGIKCVANINNECPAQLRAPGGCKDPCNVFKTDMYCCNSGSCPTEFSKFFKQRCP

DAYSYPKDDPTSTFTCPGGTNYRVVFCP

Figure 15. A) Translation product, 223 amino acids from identified open reading frame using LD139544, region 29612 to 30283. Circled amino acids identify the amino acids that form the REDD acidic cleft found in TLPs. B) PBLAST analysis identified the amino acid

sequence as a TLP specific to the plant subfamily. Triangles point to the relative positions of REDD amino acids. Lower bars indicate amino acid sequence regions that identify TLP as plant specific.

In silico analysis of the LD139544 sequence upstream of the CDS using PLACE identified core promoter and several cis regulatory elements positioned within a spatial frame work consistent with known functional genes. Elements included a TATA box 70 bp from the ATG translational start site, CA dimer consistent with transcriptional initiation 31 bases from the first T of the

TATA box, GC skew region of 41 bp encompassing the transcriptional start site (TSS), and two

CAT box sites 20 to 50 bp upstream of the TATA box (figure 16).

ATGCTACACGTTTAATTTGTTGATTAGTAGAGATTGAAGTCTCTCCTATACAATATACAT AGTCGTGTCTCAATAGTGAATTTCTACACTACGTAGTCAATACCAAGACAAGCTGTGTGT

TTATAAATA CCCTGCCAAAACTTGATCATAGTCACACCATTATTACTTGC ATGGTAGCCT AAGCCTCTCTATTTCGTAAA ATG

Figure 16. Sequence 200 bp upstream of the TLP ATG translation start site (circled). Grey highlights outline two CAAT BOXs 20 to 50 bp upstream of the TATA box (bold font). Boxed region indicates GC skew region of 41 bp encompassing the transcriptional start site (bold italic font).

Plant specific TLP isoforms.

The predicted CDS for TLP from LD139544 was used as query sequence in a BLAST search of the hop reference genome assemblies to determine if other TLP alleles or isoforms were present.

Several scaffolds showed some sequence similarity to the TLP CDS (table 6).

Acession Scaffold Size (bp) Region (bp) LD134208 84970 71896 to 72778 LD134208 84970 71197 to 72000 LD135943 69298 38250 to 39000 LD135943 69298 44805 to 45614 LD135943 69298 45800 to 46700 LD137589 85301 35067 to 35473 LD144333 83170 8347 to 10060 LD144333 83170 17303 to 17983

LD144333 83170 11289 to 11967 LD147884 36660 33854 to 34650 LD159613 24342 11737 to 12533 LD165398 20268 14669 to 15485 LD168796 18076 10500 to 11700 LD174925 14760 342 to 1003 LD184211 10994 9576 to 10255 LD184638 10807 8058 to 8906

Table 6. List of scaffolds with regions containing sequence similarity to TLP CDS. Ten complete

plant specific TLP CDS were detectable in the above scaffolds with bold font.

LD134208 71197 to 72000 MVLYLSFASTHTETLTRFNITNNCSFDVWAAAMPGGGQHLSPGQTWGLDVISGTEG RIWARTGCVFNSTGQGRCASGDCDGVLECQSKGRAPHTLAEFSLN KVKGNDFLDISLIEGFNIPMEISPSSTKQCDRRVKCAADINGPCPMELRDPEGCNNPCTVFGNDQFCCRSGGCEPTSYSKYFKDHCPDVYTYPQDDDSTSSYTCP NGTNYKVVFCP

LD134208 71896 to 72778 MRSQSIFSFLLALLYFSNSAHTIRFYITNNCPHTVWAAAIPGGGRQLISGETWILDVNQSTKGARIWARTGCRFDEDGRGKCDTGDCGGVLDCQIGGQPPKTM AEYALTQTNNLDFFDISLVDGFNVPMEFSPTSLNVHVDYSPTALCTKGIKCVVNITKDCPAELRAPGGCHNPCTVFKTAEYCCTSGRCGPTDYSRFFKERCPDAY SYSLDDQTSTFTCPAWINYNVVFCPSGE

LD135943 38250 to 39000 MRSQSIFSFLLALLYFSNSAHTIRFYIINNCPHTVWAAAIPGGGRQLISGETWILDVNQSTKGARIWARTGCRFDEDGRGKCDTGDCGGVLDCQSYGQPPNTLA EYVLTQDNKLEFFDISLVEGFNVPMEFSPTLLGAHVDYSIVAMCTTGIKCVANITKECPAELRAATGGCHNPCTVFKTAEYCCTSGSCGPTGYSRFFKERCPDANS YPKDDATLSFTCYGWINYNVVFCPSGE

LD135943 44805 to 45614 MVLYLSFASTHTETLTRFNITNNCSFDVWAAAMPGGGQHLSPGQTWGLDVISGTEGRIWARTGCVFNSTGQGRCASGDCDGVLECQSKGRAPHTLAEFSLN KVKGNDFLDISLIEGFNIPMEISPSSTKQCDRRVKCAADINGLCPMELRDPAGCNNPCTVFGNDQFCCRSGDGRCEPTSYSKYFKDHCPDVITYPRDYNSTSSYS CPNGTNYNVVFCP LD135943 45800 to 46700 MEVSKSSEVFFSKMRSQSIFSFLLALLYFSNSAHTIRFYITNNCPHTVWAAAIPGGGKELISGESSILDANQSTNSGRIWARTGCRFDEDGRGKCDTGDCGGVLN CQGGGQPPNTIAEYSLTQFNNLDFFDITLVDGFNVPMEFSPTSLNVHVDYSPTALCTKGIKCVVNITKDCPAELRAPGGCHNPCTVFKTAEYCCTSGRCGPTDY SRFFKERCPDAYSYSLDDQTSTFTCPAWINYNVVFCPSGE

LD144333 8347 to 10060 MGSSIANILFTIFIITTLLFVSSYAATFEIRNECPYTVWAAASPGGGHRLDRGQTWTLNVAAGTAMARIWGRTNCNFDGSGRGRCQTGDCGGLLQCQGWGQP PNTLAEYALNQFNNLDFIDISLVDGFNIPMDFSPTTGRCRGIRCTADINGQCPAQLRAPGGCNNPCTVFKTNEYCCTNGLGTCGPTTFSRFFKERCPDAYSYPQD DPSSTFTCPGGTNYRVVFCPRTSPRFPLEMVEGLSE

LD147884 33854 to 34650 MESITNNFFTLFITTTLFFVSSHAATFEIRNECSYTVWAAASPGGGRRLDRGQTWTLNVAAGTKMARIWGRTNCNFDGNGRGRCQTGDCGGVLQCQGWG QPPNTLAEYALNQFNNRDFIDISLVDGFNIPMDFSPTTGGCRGIRCTADINGQCPAQLRAPGGCNNPCTVFKTNEYCCTKGQGSCGPTPFSRFFKERCPDAYSY PQDDPSSTFTC

LD159613 11737 to 12533 MLSLANFSLSFLFILFTQFSIHTRAATFDIRNDCPYTVWAAASPGGGRRLDPGQAWNLWVAPGTAMARIWGRTNCNFDGSGRGRCQTGDCGSLECKGWGV PPNTLAEYALNQFGNMDFIDISVIDGFNIPMEFSPTSGACRGIRCTADINGQCPNELRTPGGCHNPCTVFKTNEYCCNNGRGSCGPTNFSKFFKTRCPDAYSYP QDDPTSTFTCPGGTNYRVVFCPRGSSHNFPLEMVDGNMSEKK

LD168796 10500 to 11700 MHCYLNLSQSPKMMKSILSFLLMLTTYFTSTHAARFDITNRCPFTVWAAAVPGGGRQLNSGQSWALDVNAGTTGARIWARTDCRFDGAGRGSCKTGDCGG TLQCQGYGQPPNTLAEYALNQYNNLDFFDISLVDGFNVPMEFGPTSPQCSRGSKCAAAINSECPSQLRANGGCNNPCTVFRTDQFCCNSGNCGPTDYSRFFK NRCPDAYSYPKDDATSMFACPGGTNYKVVFCP

LD184638 8058 to 8906 MHAFRSSCLLIFTFSHKMRSCVFSFLLMLTYFSASAHAARFDITNRCTYTVWAAAVPGGGRKLNSGESWPLDVNAGTKGARIWARTGCNFDGAGRGRCQTG DCGGILQCQGFGQPPNTLAEYALNQFQNLDFFDISLVDGFNVPMVFNPTSNCNRGITCNANINGECPAVLRASRGCNNPCTVFKTDQYCCNSGNCAPTDYSR FFKQRCPDAYSYSKDDQTSTFTCPGGTDYQVVFCPGEMDFMCF Figure 17. Amino acid sequence of different plant specific TLP isoforms present in the hop genome, identified with the scaffold number and region within the scaffold that encompasses the TLP CDS. REDD motif and conserved cysteine amino acids (C) are in large bold font.

Sequence analysis for open reading frames consistent with the correct amino acid length and/or primary structure for TLPs were detected in ten of sixteen scaffold regions (figure 17).

However, no scaffolds have enough sequence similarity to qRT-PCR primers within their TLP

CDS capable of producing a PCR product as produced by the qRT-PCR reaction used for determining the relative gene expression values listed in tables 3 and 4. Suggesting that only one TLP isoform (from LD139544) would be amplified and detected by the above qRT-PCR conditions.

'Glacier' HSVd HSVd free Gene bp Sample Sample Sample Total RPKM Sample Sample Sample Total RPKM Fold 1 2 3 1 2 3 Change

LD147884 33854 to 34650 644 0 3 0 3 0.08 0 1 2 3 0.06 0.79 LD134208 71896 to 72778 711 7 0 1 8 0.19 1 1 1 3 0.06 0.30 LD135943 38250 to 39000 714 5 11 1 17 0.39 0 0 1 1 0.02 0.05 LD135943 44805 to 45614 660 3 2 1 6 0.15 2 3 1 6 0.12 0.79 LD135943 45800 to 46700 750 3 1 0 4 0.09 1 0 0 1 0.02 0.20 LD144333 8347 to 10060 732 26 34 0 60 1.36 0 4 0 4 0.07 0.05 LD134208 71197 to 72000 654 4 3 2 9 0.23 9 7 22 38 0.76 3.33 LD159613 11737 to 12533 741 38 25 24 87 1.95 142 15 18 175 3.09 1.59 LD168796 10500 to 11700 705 1 4 0 5 0.12 3 11 22 36 0.67 5.68 LD184638 8058 to 8906 741 9 7 12 28 0.63 7 7 11 25 0.44 0.70

'Nugget' HSVd HSVd free Gene bp Sample Sample Sample Total RPKM Sample Sample Sample Total RPKM Fold 1 2 3 1 2 3 Change

LD147884 33854 to 34650 644 6 0 0 6 0.10 1 1 0 2 0.04 0.44 LD134208 71896 to 72778 711 6 8 1 15 0.23 11 0 0 11 0.22 0.96 LD135943 38250 to 39000 714 26 7 0 33 0.49 15 0 0 15 0.30 0.60 LD135943 44805 to 45614 660 6 6 1 13 0.21 8 0 3 11 0.23 1.11 LD135943 45800 to 46700 750 5 1 1 7 0.10 3 0 0 3 0.06 0.56 LD144333 8347 to 10060 732 51 30 1 82 1.20 20 0 0 20 0.38 0.32 LD134208 71197 to 72000 654 4 7 17 28 0.46 7 9 7 23 0.49 1.08 LD159613 11737 to 12533 741 28 55 7 90 1.30 16 13 11 40 0.76 0.58 LD168796 10500 to 11700 705 2 0 0 2 0.03 0 0 0 0 0.00 0.00 LD184638 8058 to 8906 741 26 26 5 57 0.82 11 2 11 24 0.45 0.55

Table 7. Relative fold change for TLP between, HSVd infected (HSVd) and mock inoculated (HSVd free) plants from ‘Glacier’ and ‘Nugget’ treatment groups. Fold changes calculated using RPKM expression values from high through-put sequencing data for reads mapped to the TLP identified regions in scaffolds listed. ‘Glacier’ shows a down regulation of three plant specific TLP isoforms (bold font) not observed in ‘Nugget’.

High through-put sequencing analysis using RPKM expression values from reads mapped to the plant specific TLP CDS regions identified in table 6 indicated a down regulation of three plant specific TLP isoforms in ‘Glacier’ in response to HSVd infection not observed in ‘Nugget’ (table

7). Additionally these plant specific TLP isoforms show a higher initial level of expression in

HSVd free plants than the other plant specific TLP isoforms identified in ‘Glacier’ and ‘Nugget’ listed table 6.

Sequence variations in the TLP CDS and regions upstream.

High through-put sequencing and alignments for each of the hop cultivars ‘Glacier’, ‘Nugget’,

‘Cascade’, ‘Columbus’, ‘Galena’, and ‘Willamette’ in the TLP CDS from LD139544 revealed several single nucleotide polymorphisms (SNP) between all hop cultivars sequenced. Relative to the hop cultivar ‘Nugget’, nineteen SNPs were observed in ‘Glacier’ (figure 18), resulting in eleven amino acid substitutions (table 8). Four of these amino acid substitutions at positions

83, 112, 142, and 173 are unique to ‘Glacier’ (figure 18). ‘Willamette’ ‘Cascade’, ‘Columbus’, and ‘Galena’ had thirteen, eleven, seven, and thirteen amino acid substitutions respectively.

None of these amino acid substitutions were unique to the respective cultivars relative to amino acid positions 83, 112, 142, and 173 (table 8). Additionally ‘Nugget’ has a unique amino acid (threonine) as opposed to proline at position 32 which is found in the other hop cultivars.

Threonine has a polar side-chain and proline has a non-polar side-chain. Furthermore the four unique amino acid substitutions observed in the cultivar ‘Glacier’ have significantly different polarities relative to the amino acids in ‘Nugget’, ‘Willamette’ ‘Cascade’, ‘Columbus’, and

‘Galena’. These include at position 83 a potentially positively charged lysine in ‘Glacier’ and a

96 potentially negatively charged glutamic acid in ‘Nugget’, ‘Willamette’, ‘Columbus’, ‘Galena’, and a polar glutamine side-chain in ‘Cascade’ (table 8). At position 142 ‘Glacier’ has a polar serine, all others have a non-polar isoleucine, and at position 173, ‘Glacier’ has a potentially positively charged lysine all others have a non-polar methionine (table 8).

G ATGAGGTCCTCTATTATTTTCTCATTTCTTTTAGTGCTAACTTACTTCTCCGCCTCAACCCATGCAGCCA

N ATGAGGTCCTCTATTATTTTCTCATTTCTTTTAGTGCTAACTTACTTCTCCGCCTCAACCCATGCAGCCA

G GATTCGACATCACAAACAGATGCCCCTTCACCGTCTGGGCAGCTGCCGTGCCCGGCGGTGGAAGGCAGCT N GATTTGATATCACAAACAGATGCACCTACACCGTCTGGGCAGCTGCCGTGCCCGGCGGTGGAAGGCAGCT

G GAACTCGGGCGAATCATGGACCCTTGACGTCAACGCAGGCACGAAAGGGGCTCGCATATGGGCTCGGACG N GAACTCGGGCCAATCATGGACCCTTGACGTGAACGCAGGCACGACAGGGGCTCGCATATGGGCTCGGACG

G GATTGTAATTTTGATGGGGCTGGACGCGGCAGTTGCAAGACTGGCGACTGCGGCGGCATTCTCCAGTGCC N GATTGTAATTTTGATGGGGCTGGACGCGGCAGGTGCGAGACTGGCGACTGCGGCGGCATTCTCCAGTGCC

G AAGCCTATGGGCAGCCGCCCAACACGTTGGCCGAGTACGCACTGAACCAATTCCAGAACTTAGATTTCTT N AAGCCTATGGGCAGCCGCCCAACACGTTGGCCGAGTACGCACTGAACCAATTCAAGAACTTAGATTTCTT

G CGATATCTCACTGGTGGATGGGTTCAATGTCCCCATGGACTTCAGTCCCACATCACCCCAGTGCAGCCGA Figu N reCGA 17C.ATCTC NineteenGCTGGTGGATGGGTTCAATGTCCCCATGGACTTCAGTCCCAC single nucleotide polymorphisms (boxed bold fontTTCACCCCAGTGCAGCCGA) in ‘Glacier’ (G) thaumatin like protein coding sequence relative to the hop cultivar ‘Nugget’(N). G GGGAGCAAGTGTGTGGCAAATATAAATAACGAATGCCCTGCTCAGTTGAGGGCCCCTGGAGGCTGCAAGG N GGGATCAAGTGTGTGGCAAATATAAATAACGAATGCCCTGCTCAGTTGAGGGCCCCTGGAGGCTGCAAGG

G ATCCATGTAATGTCTTTAAAACCGATAAGTATTGTTGTAACTCAGGTAACTGTGGACCCACAGATTATTC N ATCCATGTAATGTCTTTAAAACTGATATGTATTGTTGTAACTCAGGTAGCTGTGGACCCACAGATTATTC

G GAGGTTCTTTAAGCAGCGATGCCCTGATGCTTATAGTTACCCTAAGGATGATCAAACCAGCACATTTACG NGAGGTTCTTTAAGCAGAGATGCCCTGATGCTTATAGTTACCCTAAGGATGATCAAACCAGCACATTTACG

G TGCCCTGGAGGGACTAACTATAGGGTTGTCTTCTGCCCTTG N TGCCCTGGAGGGACTAACTATAAGGTTGTCTTCTGCCCTTG

Figure 18. Nineteen single nucleotide polymorphisms (boxed bold font) in ‘Glacier’ (G) thaumatin like protein coding sequence relative to the hop cultivar ‘Nugget’(N).

20 60 N MRSSIIFSFLLVLTYFSASTHAARFDIT NRCTYTVWA AAVPGGGRQLNSGQSWTLDVNAGTTGA RIWART

G MRSSIIFSFLLVLTYFSASTHAARFDIT NRCPFTVWA AAVPGGGRQLNSGESWTLDVNAGTKGA RIWART CAMRSSIIFSFLLVLTYFSASTHAARFDIT NRCPFTVWA AAVPGGGRQLNSGQSWPLDVNAGTKGA RIWART CL MRSSIIFSFLLVLTYFSASTHAARFDIT NRCPYTVWA AAVPGGGRQLNSGQSWALDVNAGTTGA RIWART GLMRSSIIFSFLLVLTYFSASTHAARFDIT NRCPYTVWA AAVPGGGRQLNSGETWPLDVNAGTKGA RIWART W MRSSIIFSFLLVLTYFSASTHAARFDIT NRCPFTVWA AAVPGGGRQLNSGQSWTLDVNQGTTGA RIWART

120

100 80 N DCNFDGAGRGRCE TGDCGG ILQCQGYGQ PPNTLAE YALNQFKNLDFF DISLVDGFN VPMDFSPTSPQ G DCNFDGAGRGSCK TGDCGG ILQCQGYGQ PPNTLAE YALNQFQNLDFF DISLVDGFN VPMDFSPTSPQ CA DCNFDGAGRGSCQ TGDCGG ILQCQGYGQ PPNTLAE YALNQFNNLDFF DISLVDGFN VPMDFSPTSPQ CL DCKFDGAGRGSC E TGDCGG ILQCQGYGQ PPNTLAE YALNQFKNLDFF DISLVDGFN VPMDFSPTSPQ GL DCNFDGAGRGRCE TGDCGG VLQCQAYGQ PPNTLAE YALNQFKNLDFF DISLVDGFN VPMDFSPTSPQ W GCKFDGAGRGRCE TGDCGG VLQCQGYGQ PPNTLAE YALNQFNNLDFF DISLVDGFN VPMDFSPTSPQ

140 160

180

200 N CSRGIKCVANINNECPAQLRAPGGCKDPCNVFKTDMYCCNSGSCGPTDYSRFFKQRCPDAYSYPKDDQ G CSRGSKCVANINNECPAQLRAPGGCKDPCNVFKTDKYCCNSGNCGPTDYSRFFKQRCPDAYSYPKDDQ CACSRGIKCVANINNECPAQLRAPGGCKDPCNVFKTDMYCCNSGSCGPTEFSKFFKQRCPDAYSYPKDDP CL CSRGIKCVANINNECPAQLRAPGGCKDPCNVFKTDMYCCNSGNCGPTDYSRFFKQRCPDAYSYPKDDP GL CSRGIKCVANINNECPAQLRAPGGCKDPCNVFKTDMYCCNSGNCGPTEFSKFFKQRCPDAYSYPKDDP W CSRGIKCVANINNECPAQLRAPGGCKDPCNVFKTDMYCCNSGSCGPTEFSKFFKQRCPDAYSYPKDDA

N TSTFTCPGGTNYKV V FCP G TSTFTCPGGTNYRV V FCP CATSTFTCPGGTNYKV V FCP CL TSTFTCPGGTNYRV V FCP GLTSTFTCPGGTNYRV V FCP WTSMFACPGGTNYKV V FCP

Figure 19. Thaumatin like protein isoform LD139544 showing protein primary structure amino acid variations between hop cultivars ‘Nugget’(N), ‘Glacier’ (G), ‘Cascade’ (CA), ‘Columbus’ (CL), ‘Galena’ (GL), and ‘Willamette’ (W). Conserved regions are boxed. Stars ( ) indicate positions of amino acids (circled, ) unique to ‘Glacier’. Arrow ( ) indicates position of amino acid (boxed, ) unique to ‘Nugget’. ( ) indicate

amino acid positions that form the REDD motif. Bolded C conserved cysteines.

None of the amino acid substitutions in ‘Glacier’ involve conserved cysteine residues, regions, or members of the REDD motif within the TLP CDS recognized by Petre, et al. (2011) who identified six conserved regions, fifteen conserved cysteine residues, and an REDD motif important in formation of an acidic cleft associated with TLP antifungal activity (figure 19). However, the unique threonine at position 32 in ‘Nugget’ is within the first conserved region.

Table 8. Amino acid substitutions in ‘Glacier’ thaumatin like protein isoform LD139544 relative to ‘Nugget’ including amino acids at these positions for the hop cultivars ‘Galena’, ‘Cascade’, ‘Columbus’, and ‘Willamette’.

No sequence variations in ‘Glacier’, ‘Nugget’, ‘Cascade’, ‘Columbus’, ‘Galena’, and ‘Willamette’ are present in the immediate 100 bp upstream region of the TLP isoform LD139544 CDS which includes the core promoter region consisting of the ATG translation start site, TATA box and a

GC skew region of 41 bp encompassing the transcriptional CA start site. Sixteen SNP variations and one dinucleotide substiution exist between ‘Glacier’ and ‘Nugget’ from position -505 to -

108 additionally, in this region there exist, in ‘Nugget’, one dinucleotide and one trinucleotide insertion (figure 20).

-505 -446 N CATAATAATTAGAAAATTAGGGGCCAGCTAAATATAGGGATTCATTATTTGCTTAGTTTT G CATAATAATTAGAAAATTAGGGGCCAGCTAAATATAGGGATTCATTATTTACTTAGTTTT -445 -386 N TATATTTATTAGTGTGCATGGCATAATTAAGGGAAAAAGTAAAAACAGAAAGATTTAAGT G TATATTTATTAGTGTGCATGGCATAATTAAGGGAAAAAGTAAAAACAGAAAGATTTAAGC

-385 -326 N CACAAGATTTTATGTATAATGAATATTGCTAAGGAATATTATTAGATTATTACATATCAT G CACAAGATTTTATGTATAATGAATATTGCTAT GGAATATTATTACATTATTACACATCAT

-325 -266 N ATTTTTATATGTGTAATCACATATTTTAGTTTCAAATAAAATATGTGGGATCCGATACTA G ATTTTTAT----GTGTAATCACATATTTTAGTTTCAAATAAAATATG-----GATCCGATACTA

-265 -206 N AATCACCATAT TATATATGTTGTTAGCATGATTTTCATA AATAATGGTCATATCTAATAG G AATCACCGCAGTATATGTGTTGTTAGCATAATTTTCATGTAATAATGGTCATATCTAATAG -205 -146 N ATAGAATAATGCTACACGTTTAATTTGTTGATTAGTAGAGATTGAAGTCTCCTATACAAT G C TAGAATAATGTTAT ACGTTTAATTTGTTGATTAGTAGAGATTGAAGTCTCCTATACAAT

-145 -86 N ATACATAGTCGTGTGTCAATAGTGGATTTCTACACTGCGTAGTCAATACCAAGACAAGCT G ATACAAAGTCGTGTGTCAATAGTGGATTTCTACACTACGTAGTCAATACCAAGACAAGCT

-85 -27 N GTGTTTT TATAAATA CCCTGCCAAAACTTGATCATAGT CA ACCATTATTACTTGCATGG G GTGTTTT TATAAATA CCCTGCCAAAACTTGATCATAGT CA ACCATTATTACTTGCATGG -26 +1 N TAGCCTAAGCCTCTCTATTTCGTAAAATG G TAGCCTAAGCCTCTCTATTTCGTAAAATG

Figure 20. Positions of single nucleotide substitutions and multi nucleotide insertions (bold type in ovals) and position of TATA box and CA transcriptional start (thick boxes) within the 41 bp GC rich region (box shaded light grey without lined borders). Cis-regulatory elements unique to ‘Glacier’ (G) and ‘Nugget’ (N) are marked with light and dark grey outlined boxes, respectively.

100

In total 23 nucleotide variations exist between ‘Glacier’ and ‘Nugget’ in the 397 bp region up stream of the first CAAT cis regulatory element (CRE). These variations create eleven unique

CREs, 6 in ‘Nugget’ not present in ‘Glacier’ and 5 in ‘Glacier’ not present in ‘Nugget’. However the two CAAT CREs within the core promoter region are conserved between these two hop cultivars (figure 20). In addition 102 CREs, which are conserved between the two cultivars have been identified (table 9).

101

Position in 'Glacier' Position in 'Glacier' 'Glacier' and 'Nugget' with (+) or (-) strand 'Glacier' and 'Nugget' with (+) or (-) strand indication indication HDZIP2ATATHB2 3 (+) TAATMATTA POLASIG1 213 (+) AATAAA POLASIG3 4 (+) AATAAT LECPLEACS2 199 (-) TAAAATAT POLLEN1LELAT52 11 (+) AGAAA ROOTMOTIFTAPOX1 218 (-) ATATT GT1CONSENSUS 12 (+) GRWAAW GATABOX 229 (+) GATA SORLIP2AT 21 (+) GGGCC CACTFTPPCA1 231 (+) YACT CARGCW8GAT 28 (+) CWWWWWWWWG ARR1AT 236 (-) NGATT ROOTMOTIFTAPOX1 31 (-) ATATT GTGANTG10 238 (-) GTGA ARR1AT 38 (+) NGATT POLASIG3 276 (+) AATAAT POLASIG3 44 (-) AATAAT WBOXNTERF3 282 (-) TGACY TATABOX5 45 (+) TTATTT WRKY71OS 283 (-) TGAC SEF4MOTIFGM7S 56 (+) RTTTTTR GATABOX 287 (-) GATA MARARS 58 (+) WTTTATRTTTW CPBCSPOR 290 (-) TATTAG SEF1MOTIF 62 (+) ATATTTAWW 10PEHVPSBD 299 (-) TATTCT ROOTMOTIFTAPOX1 62 (+) ATATT POLASIG3 301 (+) AATAAT POLASIG1 65 (-) AATAAA ACGTATERD1 312 (-) ACGT CPBCSPOR 67 (+) TATTAG ACGTATERD1 312 (+) ACGT CACTFTPPCA1 71 (-) YACT POLASIG2 315 (-) AATTAAA RYREPEATLEGUMINBOX 74 (-) CATGCAY RAV1AAT 322 (-) CAACA RYREPEATBNNAPA 75 (-) CATGCA ARR1AT 325 (+) NGATT CARGCW8GAT 82 (-) CWWWWWWWWG CACTFTPPCA1 330 (-) YACT CARGCW8GAT 82 (+) CWWWWWWWWG INRNTPSADB 335 (-) YTCANTYY GT1CONSENSUS 92 (+) GRWAAW ARR1AT 335 (+) NGATT GT1CONSENSUS 93 (+) GRWAAW CAATBOX1 337 (-) CAAT GT1GMSCAM4 93 (+) GAAAAA SURECOREATSULTR11 343 (-) GAGAC DOFCOREZM 96 (+) AAAG CAATBOX1 353 (+) CAAT CACTFTPPCA1 98 (-) YACT ROOTMOTIFTAPOX1 354 (-) ATATT SEF4MOTIFGM7S 100 (-) RTTTTTR DPBFCOREDCDC3 366 (-) ACACNNG POLLEN1LELAT52 107 (+) AGAAA BIHD1OS 370 (+) TGTCA DOFCOREZM 109 (+) AAAG WBOXATNPR1 371 (-) TTGAC NODCON1GM 109 (+) AAAGAT WRKY71OS 371 (-) TGAC OSE1ROOTNODULE 109 (+) AAAGAT CAATBOX1 373 (+) CAAT ARR1AT 111 (+) NGATT NRRBNEXTA 376 (+) TAGTGGAT ARR1AT 125 (+) NGATT CACTFTPPCA1 377 (-) YACT ROOTMOTIFTAPOX1 142 (-) ATATT ARR1AT 380 (+) NGATT ROOTMOTIFTAPOX1 143 (+) ATATT POLLEN1LELAT52 383 (-) AGAAA CAATBOX1 145 (-) CAAT CACTFTPPCA1 389 (+) YACT ROOTMOTIFTAPOX1 155 (-) ATATT WBOXHVISO1 397 (-) TGACT ROOTMOTIFTAPOX1 156 (+) ATATT WBOXNTERF3 397 (-) TGACY POLASIG3 158 (-) AATAAT WBOXATNPR1 398 (-) TTGAC POLASIG3 166 (-) AATAAT WRKY71OS 398 (-) TGAC NAPINMOTIFBN 171 (+) TACACAT CAATBOX1 400 (+) CAAT ROOTMOTIFTAPOX1 179 (+) ATATT SEF4MOTIFGM7S 419 (+) RTTTTTR SEF4MOTIFGM7S 181 (+) RTTTTTR TATABOX2 424 (+) TATAAAT NAPINMOTIFBN 187 (-) TACACAT WBOXHVISO1 452 (-) TGACT ARR1AT 193 (-) NGATT WBOXNTERF3 452 (-) TGACY GTGANTG10 195 (-) GTGA WRKY71OS 453 (-) TGAC CARGCW8GAT 198 (-) CWWWWWWWWG GTGANTG10 454 (-) GTGA CARGCW8GAT 198 (+) CWWWWWWWWG POLASIG3 461 (-) AATAAT LECPLEACS2 199 (-) TAAAATAT CACTFTPPCA1 466 (+) YACT ROOTMOTIFTAPOX1 199 (+) ATATT RYREPEATBNNAPA 470 (-) CATGCA TATABOX5 212 (-) TTATTT S1FBOXSORPS1L21 473 (+) ATGGTA

Table 9. List of cis-regulatory elements common to ‘Glacier’ and ‘Nugget’.

102

Survey for the presence of HSVd by RT-PCR in Washington State.

The three major hop growing regions in Washington State include the Lower Yakima Valley,

Moxee and Toppenish Creek. HSVd was identified in each of these regions. Rates of HSVd infection varied between 9.6 to 72.3 % and were cultivar dependent (table 10). The overall average of HSVd infection was 17.34 % for the 1,635 samples tested. ‘Galena’, Glacier’,

‘Cascade’, ‘Tomahawk’, and ‘Nugget’ had the highest incidence at 72.3, 56.3, 31.3, 24.5 and

21.1 %, respectively. One cultivar, ‘Palisade’ did not test positive for HSVd. However, the survey sample size for this cultivar was limited to 15.

Cultivar ID Total samples tested Samples tested positive Percent positive

Apollo 176 28 15.9 Bravo 10 2 20 Cascade 195 61 31.3 Centennial 188 18 9.6 Chinook 231 30 13 Columbus 31 4 12.9 Exp Cluster 15 1 6.7 Galena 47 34 72.3 Glacier 16 9 56.3 Nugget 57 12 21.1 Palisade 15 0 0 Simcoe 65 6 9.2 Sterling 16 1 6.3 Super Alpha 65 5 7.7 Super Galena 126 14 11.1 Tomahawk 98 24 24.5 Unidentified VGXPOI 73 13 17.8 Vanguard 32 3 9.4 Zeus 179 20 11.2 Total 1635 285 17.34

Table 10. Survey results for the presence of HSVd separated by cultivar from the three major hop growing regions in Washington State.

103

DISCUSSION

Hop stunt disease, now known to be caused by Hop stunt viroid, was first recognized in Japan over 70 years ago, and reached epidemic proportions in that country two decades following its discovery. HSVd has now been detected worldwide and genetically variable isolates have been identified in a variety of agriculture crops including grapevine, citrus, stone fruits, and hop. Hop stunt disease was first detected in hop gardens in Washington State in 2004, sixty years after being discovered in Japan. The association with HSVd in WA was subsequently confirmed. A

2012 survey for the presence of HSVd by RT-PCR, with leaf samples collected from the three major hop growing regions in Washington State indicated an overall incidence of 17.34% ranging from 6.3 to 72.3% depending on cultivar. One thousand six hundred and thirty-five leaf samples were collected on FTA cards from 20 hop cultivars. ‘Galena’, Glacier’, ‘Cascade’,

‘Tomahawk’, and ‘Nugget’ had the highest incidence at 72.3, 56.3, and 31.3, 24.5 and 21.1% respectively. These results are consistent with a limited 2004 survey (Eastwell and Nelson,

2007) of 126 hop plants, representing 12 known hop cultivars and 9 unidentified hop cultivars sampled from the same three growing regions. Results from the 2004 survey indicated an overall incidence of 15.1%. The slight change in overall incidence of HSVd infected hop plants from the limited 2004 survey to the much larger 2012 survey suggest that the spread of HSVd is slow, but occurring in spite of the increased use of viroid-tested planting stock. There can be a cultivar-dependent latent period between infection with HSVd (Kawaguchi-Ito et al., 2009) and the first appearance of visible symptoms, making it hard to determine the current economic impact. However, as the latent period for currently infected plants is reached one can expect to

104 see an increasing economic effect. In addition, it has been reported that hop gardens with high incidences of HSVd have significantly higher incidences of fusarium canker wilt disease, the increased presence of this disease is independent of cultivar tolerances to HSVd (Ocamb, 2016).

Mixed infections such as this where infection of one pathogen, such as HSVd increases the incidence or severity of another disease worsens the economic impact. Mixed infections which sensitize hop to another pathogen highlight the need to limit the spread of HSVd, regardless of the use of HSVd tolerant cultivars.

A five year study comparing HSVd infected and uninfected hop cultivars ‘Glacier’, ‘Nugget’,

‘Cascade’, ‘Columbus’, ‘Galena’, and ‘Willamette’ for alpha and beta acid production, cone yield, and plant growth revealed ‘Nugget’ and ‘Galena’ to be the most tolerant hop cultivars to

HSVd infection and ‘Glacier’ to be the most severely affected (Eastwell, unpublished result).

Data from the aforementioned studies were used as the basis for choosing hop cultivars for this study. Whole genome sequencing and RNAseq analysis were coupled with high through-put sequencing to identify potential genes involved in the molecular mechanisms associated with hop stunt disease and to identify new DNA-markers that distinguish HSVd-tolerant plants from

HSVd-sensitive plants.

HSVd infected ‘Glacier’ showed reduced growth over the same time period when compared to healthy ‘Glacier ‘and ‘Nugget’, and HSVd infected ‘Nugget’. These observations were consistent with the above mentioned five year field trial. Leaves from HSVd or mock inoculated plants

105 paired for similar size, age, and growing location on the plant were sampled at various times from 30 to 270 days post inoculation as a tissue source for RNA extraction. RPKM expression values from RNAseq analysis of RNA samples from healthy and HSVd infected ‘Glacier’ and

‘Nugget’ revealed different gene expression profiles between cultivars, and differences between HSVd infected and healthy plants of the same cultivar. One isoform (LD139544) of thaumatin like protein (TLP), a pathogenesis related protein, was observed to be down regulated 12-fold in ‘Glacier’ bines infected with HSVd compared to HSVd-free ‘Glacier’; no change in this isoform of thaumatin like protein expression was observed in ‘Nugget’ as a consequence of HSVd infection. Exploratory RNAseq results were confirmed using qRT-PCR.

However, the down regulation (2.6) was lower than that calculated using RPKM expression values. Differences between ‘Nugget’ treatment groups showed no statistical significance The differences in fold change values for TLP expression observed between RNAseq RPKM expression values and qRT-PCR results is not unexpected for several reasons. Sample size used to generate RPKM values was low, consisting of three samples for each treatment subgroup compared to 20 or greater samples used to generate qRT-PCR expression values. RPKM values are not normalized to an internal control to account for potential discrepancies in starting RNA concentration or discrepancies that result from the RNA library processing steps that can bias the relative abundance of some RNA transcripts and thus affect RNAseq RPKM values.

Additionally, the detection methods for measuring transcript abundance using RNAseq and qRT-PCR are fundamentally different, discrete digital measurements are used to calculate

RPKM values versus continuous analog signals used to quantify qRT-PCR values.

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Differences in gene expression patterns as observed for different TLP isoforms, including

LD139544, between HSVd infected ‘Glacier’ and ‘Nugget’ can be the result of epigenetic differences due to different DNA methylation patterns between cultivars or individual bines, or genetic variations in the DNA sequence of the promoter regions between the two cultivars, or differences in the synthesis of regulatory factors. Genetic variations in the DNA sequence between cultivars can be identified by whole genome sequencing. Sequencing of the TLP isoform LD139544 CDS, core promoter region and proximal CREs from the HSVd sensitive cultivar ‘Glacier’ and cultivars with varying degrees of tolerance to HSVd infection, including

‘Nugget’ was done to identify genetic markers which would distinguish the HSVd sensitive cultivar from HSVd tolerant cultivars.

Whole genome sequencing of DNA samples extracted from leaf tissue taken from hop cultivars

‘Glacier’, ‘Nugget’, ‘Cascade’, ‘Columbus’, ‘Galena’, and ‘Willamette’ was done by high through- put sequencing. Alignments of the whole genome sequencing results revealed no sequence variations in the immediate 100 bp upstream region of the TLP 139544 isoform CDS which includes the core promoter region consisting of the ATG translation start site, TATA box and a

GC skew region of 41 bp encompassing the transcriptional CA start site. However, sixteen SNP variations and one dinucleotide substiution exist between ‘Glacier’ and ‘Nugget’ from position -

503 to -106. Additionally, in this region in ‘Nugget’, there exists one dinucleotide and one trinucleotide insertion relative to ‘Glacier’. In total, 23 nucleotide variations exist between

‘Glacier’ and ‘Nugget’ in the 397 bp region up stream of the first CAAT cis-regulatory element.

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These variations create eleven unique CREs, 6 in ‘Nugget’ not present in ‘Glacier’ and 5 in

‘Glacier’ not present in ‘Nugget’. Within the LD139544 TLP CDS, SNPs were observed in

‘Glacier’ relative to ‘Nugget’. ‘Willamette’, ‘Cascade’, ‘Columbus’, and ‘Galena’ with nineteen, thirteen, eleven, seven, and thirteen substitutions, respectively. Several of these SNPs result in amino acid substitutions. Four of the SNPs within the ‘Glacier’ TLP CDS result in amino acid substitutions unique to ‘Glacier’. ‘Nugget’, the cultivar shown to be most tolerant to HSVd in field studies, has a unique SNP which results in a codon for the amino acid (threonine) at position 32 as opposed to proline, which is found at this position in the other hop cultivars sequenced. In total 42 SNPs were identified in TLP isoform LD139544 that can be used in marker assisted breeding programs to distinguish potential HSVd tolerant plants from HSVd sensitive plants.

Consequences of protein primary sequence variations listed above among hop TLPs and reduced TLP isoform LD139544 levels in ‘Glacier’ are potentially multifaceted. The presence of a conserved REDD motif in each of the hop TLPs sequenced has been associated with antifungal activity (Petre, et al., 2011). However, TLP antifungal activity has been shown to involve several different mechanisms which can lead to fungal plasma membrane disruption (Liu et. al., 2010).

Specific substrate or ligand binding has not been precisely correlated to a specific TLP primary structure or antifungal activity. Several TLPs with antifungal activity have been shown to have one or more of the following properties: -1,3 glucan binding, endo--1,3-glucanase, and endo-

-1,4-xylanse inhibition (Liu et. al., 2010). However, Van Loon et al. (2006) demonstrated that

108 having one of the above listed properties does not guarantee that a specific TLP will have antifungal activity. Fierens et. al. (2007) demonstrated that the binding properties of individual

TLPs with a specific antifungal property, such as, endo--1,4-xylanase inhibition, are specific to both the TLP primary structure and the substrate primary structure; e.g. TLXI a TLP isolated from wheat is capable of inhibiting a endo--1,4-xylanase XynI from Tricoderma longibrachiatum but not endo--1,4-xylanase XynII. Such specificities are not unique to TLPs, but are a fundamental principal between substrate/enzyme and ligand/receptor protein interactions. For ‘Glacier’, a down regulation in TLP expression as a result of HSVd infection could result in an increased susceptibility to fungal infection which would not be experienced by a similarly infected ‘Nugget’. It has been reported by Ocamb (2016) that hop gardens with high incidences of HSVd have significantly higher incidences of fusarium canker, which is caused by the fungal pathogen Fusarium sambucinum, and the presence of this disease may be an indicator for the presence of HSVd even in HSVd-tolerant hop cultivars, such as ‘Nugget’.

Additionally, Narasimhan et. al. (2005) demonstrated that the TLP protein osmotin from tobacco binds a Saccharomyces cerevisiae G protein-coupled receptor that is important in the signal cascade that regulates lipid and phosphate metabolism. G protein-coupled receptors

(GPCR) are a large class of transmembrane receptors that activate a wide range of cellular responses involved in plant growth, development, and resistance to abiotic and biotic stress. If

TLPs in hops, including ‘Glacier’ and ‘Nugget’ behave as ligands to (GPCR), the potential for altered phenotypes resulting from disruption of a signal cascade, as a result of down regulated

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TLP expression exists. Such changes could include increased sensitivity to pathogens, diminished plant growth, altered development, reductions in secondary metabolites, such as alpha and beta acids as seen in HSVd infected ‘Glacier’, and reduced or varied flowering as observed in the HSVd sensitive cucumber Cucumis sativus L. cv. Suyo Long.

Variations in gene expression patterns that result in the various manifestations of hop stunt disease symptom severity (whether a result of a signal cascade event that regulates the synthesis of specific proteins such as TFs which bind CREs, DNA sequence variations in host promoters that are capable of binding different TFs, or a change in the epigenetic profile) must be kept in context with the genotype of the infecting HSVd isolate, which also influences symptoms expressed by hop cultivars. Because symptom severity is dependent on the genotype of the HSVd isolate, this study utilized a single HSVd isolate cloned and maintained in plasmid pCR2.1 for use as an inoculum. Prior to inoculating plants the sequence of the HSVd isolate was determined to be identical to HSVd genotype hKFKi (GenBank GI:12082110). Isolation and sequencing of HSVd recovered from infected study plants was verified to be unchanged during the course of the study. Additionally, changes in gene expression patterns can be influenced by age, tissue type, and abiotic and biotic stresses related to growing conditions and exposure to insects and/or other plant pathogens. Each of these potential confounders were mitigated by using hop plants of the same age, maintained under identical growing conditions, tested and shown to be free of virus and viroid pathogens (with the exception of hop latent viroid), and continually monitored for insects, other plant pathogens.

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Several models have been proposed to describe the mechanisms by which viroids induce disease symptoms. Because viroids lack coding RNA, any model requires that the viroid RNA must directly interact with either a host protein, nucleic acid, or both. Since viroids do not code for proteins they must co-opt the use of host RNA polymerases for replication. One hypothesis is the act of diverting host enzymes to replicate viroids causes a general disruption

(attenuation) in the production of host RNAs, and disease symptoms would be a side effect of this disruption. However, latent viroids such as Hop latent viroid are capable of infecting several hop cultivars without expression of diseases symptoms (Barbara et. al., 1990). Additionally, it has been shown that viroids are capable of binding proteins other than host RNA polymerases.

Gozmanova et al. (2003) identified a bromodomain-containing protein from tomato that was used for nuclear transport by PSTV and HSVd has been shown to bind cucumber phloem protein 2 protein as a possible chaperon in long distance transport (Gomez and Pallas, 2001;

Owens et al., 2001). Two kinase proteins involved in signal cascades have also been shown to be bound by viroids. Recently, Wang et al. (2014) showed that PSTV is capable of binding transcription factor TFIIIA. Binding of such proteins by viroids could cause a diversion of proteins from their intended physiological role leading to disease symptom expression.

Whereas the general down regulation of genes predicted by the redirected use of RNA polymerase to replicate viroids, the consequences of viroid binding to specific proteins would lead to targeted disruptions of specific molecular pathways in the host. Such specificity would account for observed differences in the types and severity of disease symptoms expressed between tolerant, sensitive, and resistant cultivars as a result of host genotype differences.

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Other models which account for differing disease expression patterns associated with viroid and host genotype differences are the different RNA silencing mechanisms triggered by double- stranded RNA such as replicative intermediates associated with viroid replication and single- stranded RNA that has double-stranded secondary structure similar to that of HSVd and other members of the Pospiviroidae family. RNA silencing mechanisms include TGS involving sRNA directed DNA methylation of host DNA and PTGS involving sRNA directed degradation of host encoded RNA transcripts and translation repression. Direct evidence supporting RNA silencing as a mechanism by which viroids or viruses induce disease symptoms as a result of infection has recently emerged. Shimura et al. (2011) and Smith et al. (2011) independently showed that leaf yellowing in Nicotiana tabacum following infection with CMV and its Y-satellite RNA is directly caused as a result of PTGS. The pathogenic determinant was mapped to a 24 bp fragment within the 349 bp Y-satellite RNA that is used as a template for the production of a 22 bp sRNA which targets (that is, it is complementary to) a specific 22 bp sequence in the host CHLI mRNA directing its cleavage. CHLI is an important gene in the host chlorophyll biosynthetic pathway.

Similar infections with CMV and its Y-satellite RNA in A. thaliana and tomato did not result in leaf yellowing. However, site directed mutagenesis of the Y-satellite RNA which changed its sequence to match that of the CHLI mRNA in A. thaliana and tomato resulted in the production of small RNAs and subsequent PTGS of the CHLI mRNA, leading to a leaf yellowing phenotype in both A. thaliana and tomato. Additional direct evidence for sRNA induced symptoms has been reported by Navarro et al. (2012) who demonstrated that peach calico disease is caused by specific genotypic isolates of PLMVd that have an explicit 12 to 14 bp insertion. As a result of

112 this insertion, two 21 bp viroid-derived sRNAs (PC-sRNA8a and b), which differ by one nucleotide position, are produced and are complementary to a 21 bp sequence of peach mRNA encoding for a chloroplastic heat-shock protein (cHSP90) important in chloroplast development.

Furthermore, it was demonstrated the cHSP90 mRNA was cleaved at predicted positions based on sequence homology between cHSP90 and the two viroid derived small RNAs. PLMVd isolates lacking the 12 to 14 bp insertion which are necessary for creating the complementary 21 bp sequence to peach cHSP90 mRNA are incapable of producing PC-sRNA8a and b, and do not induce peach calico disease. Furthermore, a report by Eamens et al. (2014) provide indirect evidence for PTGS in Nicotiana species transgeniclly expressing small RNAs complementary to the pathogenic region of PSTVd. Martinez et al. (2014) showed that infection of N. benthamiana resulted in specific alterations of DNA methylation patterns to ribosomal RNA genes consistent with TGS of these genes. The association of viroid RNAs with PTGS, TGS and direct interactions with host proteins, suggest that viroids are capable of inducing disease symptoms through several different molecular mechanisms.

Owens et al., (2012) showed that viroids are capable of inducing the production of pathogenesis proteins, and Singh (1971) showed that viroids can elicit a plant response similar to a hypersensitive response. It is well documented that pathogen invasion can induce an innate plant immune response which involves up regulated expression of several different PR proteins, including TLPs, that are produced systemically in the plant. Previous studies have shown PR proteins to be induced in several viroid-infected plants such as, tomato and potato.

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The viroids include the Pospiviroidae family viroids PSTVd, CEVd, CPFVd, and CSVd, (Conejero et al., 1979; Camacho-Henriquez and Sanger, 1982). Therefore, the observed down regulation

(versus an expected up regulation as seen in other viroid plant interactions) of the PR protein

TLP isoform LD139544 in HSVd infected ‘Glacier’ may be indicative of the ability of HSVd to attenuate the innate immune response in HSVd sensitive cultivars such as ‘Glacier’ but not in tolerant cultivars like ‘Nugget’, which show a slight up regulation of TLP isoform LD139544 upon

HSVd infection.

Interestingly RPKM expression values from RNAseq analysis of RNA samples from healthy and

HSVd infected ‘Glacier’ and ‘Nugget’ revealed that transcriptional levels for valerophenone synthase, the enzyme used to produce the first intermediate, PIVP, in the alpha acid biosynthetic pathway, was not affected in HSVd-infected ‘Glacier’ or ‘Nugget’. This is in contrast to the reductions in alpha acid production observed in the above mentioned field trial.

However, the lack of change in valerophenone synthase transcripts could be due the relatively short time study plants had been infected. Kawaguchi-Ito et al. (2009) in a 15 year study which tracked phenotypic changes in the hop cultivar ‘Kirin II’ demonstrated that severity of symptoms, including the reduction of alpha acid production, increased as the length of time plants were infected with HSVd increased. Plants infected with four different strains of HSVd

(HSVd-hop, HSVd-plum, HSVd-citrus, and HSVd-grapevine) all showed increased symptom severity with increased duration of the infection, but the relative time for increased symptom expression is different for each HSVd strain. Hops infected with HSVd-grapevine or HSVd-hop

114 did not show statistically significant stunting until year 5. Pronounced bine stunting with bent and curled leaves did not appear until year 7. Alpha acid content showed no decrease in infected cones during the first season, but diminished severely in all HSVd-infected cones by the second season, showing a reduction of between 46% to 55% in alpha acid content compared to healthy controls. Had HSVd infected plants in this study been allowed to pass through a period of dormancy and complete two growing seasons, it is likely that a reduction in alpha acid production would have been observed.

Direct evidence for small RNA mediated gene silencing has recently been demonstrated for viroid-induced mRNA degradation that is indicated as a down regulation in RPKM expression profiles similar to those observed for TLP LD139544. Attempts to create a sRNA profile by high through-put sequencing, similar to the RNA expression profile used in RNAseq analysis, were unsuccessful. However, to explore the possibility that HSVd was mimicking sRNA-mediated gene silencing of TLP directly, a library consisting of ninety-six 25 bp fragments resembling small

RNAs was created in silico from the HSVd hKFKi sequence. None of the in silico generated sRNAs aligned to the TLP LD139544 CDS or the 500 bp region directly up stream of the CDS, indicating that it is unlikely that sRNAs from the hKFKi HSVd genome are directly interacting with the TLP gene LD139544 or TLP transcripts to cause a down regulation of this TLP. However, this does not rule out the possibility that sRNA-mediated gene silencing is indirectly regulating transcript levels of this TLP via a down regulation of one of the many transcription factors that bind to

CREs located up stream of the TLP core promoter region. A detailed analysis of the CREs unique

115 to the LD139544 TLP regulatory region in ‘Glacier’ and the sequences of the various transcription factors involved in forming transcription complexes with these CREs and the CREs used to regulate each of the identified transcription factors has the potential to discover a sequence that satisfies the alignment parameters which govern the different sRNA mediated

TGS or PTGS gene silencing mechanisms. Additionally, studies that catalog differences in DNA methylation patterns between plants tolerant and sensitive to HSVd may also reveal patterns that are associated with HSVd tolerance.

Though this study did not identify a clear link of the molecular mechanisms associated with hop stunt disease, it has provided new DNA-markers that have the potential to distinguish HSVd- tolerant plants from HSVd-sensitive plants. In addition the results of this work have identified potential genes involved in regulating the viroid host interaction. Further functional studies using these identified genes are needed to establish if they are directly or indirectly involved, or independent of the molecular mechanisms associated with the HSVd plant interactions which regulate the differences in severity of expressed symptoms observed in HSVd tolerant and sensitive hop cultivars.

One of the difficulties of investigating the ability of TLPs to act as ligands to GPCR or other signal cascade receptors is the abundance of redundant TLP isoforms in model plants such as arabidopsis, which have as many as 24 TLP isoforms (Petre et. al., 2011; Deihimi, et. al., 2012). A

NCBI (BLASTn) search of the 132,476 hop reference genome scaffolds (GenBank assembly

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GCA_000831365.1) using the hop TLP CDS from LD139544 identified in this study detected ten additional plant specific TLP isoforms. However, based on sequence variations each of these isoforms expression patterns could be followed and quantified by qRT-PCR during the course of infection. The ability to quantify each isoforms expression level combined with the limited number of potential TLP isoforms relative to that of arabidopsis in hops would enhance the utility of future studies investigating TLP binding to receptors and other proteins. The use of hop in such studies would reduce, but not eliminate, the complexity of multiple TLP isoforms which have similar protein binding and biological properties and allow for a more accurate assessment of the many molecular functions associated with TLP expression patterns.

The identification of different expression patterns for one TLP isoform (LD139544) by this study in hop cultivars infected with HSVd could provide the bases for an assay that identifies current and future hop cultivars that have an increase in TLP expression of TLP isoform LD139544 and additional TLP isoforms as opposed to a decrease as seen in ‘Glacier’ or no change as observed in ‘Nugget’ when infected with HSVd. Such cultivars, because of their ability to increase their

TLP expression levels as a result of HSVd infection are likely to be more resistant to fungal pathogens such as Fusarium sambucinum, the causative agent of fusarium canker in hops.

Several studies have shown that an over-expression of TLPs and other PR5 proteins in transgenic plants have resulted in increased disease resistance to fungal pathogens, e.g. osmotin increases in potato heightens resistance to Phytophtohra infestans (Liu et al., 1994), different rice TLP isoforms boosts resistance to Rhizoctonia solani and Fusarium graminearum,

117 the causative agents of rice sheath and head blight, respectively (Datta et al., 1999);

(Mackintosh, et al. 2007), and transgenic grapes constitutively expressing the Vitis vinifera TLP

VVTL-1, display persistent resistance to several fungal pathogens such as Uncinula necator and

Botrytis cinerea (Dhekney et al., 2010). Hop cultivars that naturally increase their expression of

TLPs as a result of HSVd infection are likely to show similar resistance patterns to fungal infections as transgenic plants that overexpress TLPs. Selection of these hop cultivars for production could reduce the economic impact of fungal infections on plants that become infected with HSVd.

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REFERENCES

Allen, E., Xie, Z., Gustafson, A. M., and Carrington, J. C. 2005. MicroRNA-directed phasing during trans-acting siRNA biogenesis in plants. Cell 121:207–21.

Astruc, N., Marcos, J. F., Macquaire, G., Candresse, T., and Pallas, V. 1996. Studies on the diagnosis of hop stunt viroid in fruit trees: Identification of new hosts and application of a nucleic acid extraction procedure based on non-organic solvents. European J. Plant Pathol. 102: 837-846.

Axtell, M. J. 2013. Classification and Comparison of Small RNAs from Plants. Annu. Rev. Plant Biol. 64:137–59.

Axtell, M. J., Jan, C., Rajagopalan, R., and Bartel, D.P. 2006. A two-hit trigger for siRNA biogenesis in plants. Cell 127:565–77.

Barbara, D. J., Morton, A., Adams, A. N., and Green, C. P. 1990. Some effects of hop latent viroid on two cultivars of hop (Humulus lupulus) in the UK. Ann. Appl. Biol. 117:359-366.

Basehoar, A. D., Zanton, S.J., and Pugh, B. F. 2004. Identification and distinct regulation of yeast TATA box-containing genes. Cell. 116:699–709.

Beatson, R. A. 1993. Breeding and development of hop cultivars in New Zealand. Proceedings of the Scientific Commission of the International Hop Growers' Committee, Wye, England. 12-18.

Beatson, R. A., Ansell, K. A., and Graham, L. T. 2003. Breeding, development, and characteristics of the hop (Humulus lupulus) cultivar ‘Nelson Sauvin’. New Zeal. J. Crop. Hort. 31(4):303- 309.

Binder, S., Knill, T., and Schuster, J. 2007. Branched-chain amino acid metabolism in higher plants. Physiol. Plant. 129:68–78.

Blumenthal, T. 2004. Operons in eukaryotes. Briefings in Functional Genomics and Proteomics. (3)3:199–211.

Borsani, O., Zhu, J., Verslues, P. E., Sunkar, R., and Zhu, J. K. 2005. Endogenous siRNAs Derived from a Pair of Natural cis-Antisense Transcripts Regulate Salt Tolerance in Arabidopsis Cell. 123:1279–1291.

Boycheva, S. 2014. The rise of operon-like gene clusters in plants. Trends Plant Sci. (7):447-59.

119

Branch, A. D., Benenfield, B. J., and Robertson, H. D. 1988. Evidence for a single rolling circle in the replication of potato spindle tuber viroid. Proc. Natl. Acad. Sci. U. S. A. 85:9128– 9132.

Breathnach, R., and Chambon, P. 1981. Organization and expression of eukaryotic split genes coding for proteins. Annu. Rev. Biochem. 50:349–383.

Briggs, D. E., Boulton, C. A., Brookes, P. A., and Stevens, R. 2000. Brewing science and practice. Woodhead Publishing Limited. Cambridge.

Brodersen, P., and Voinnet, O. 2006. The diversity of RNA silencing pathways in plants. Trends in Genetics. 22:268-280.

Burley, S. K. 1996. Biochemistry and structural biology of transcription factor IID(TFIID) Annu. Rev. Biochem. 165:769-799.

Camacho-Henriquez, A., and Sanger, H. L. 1982. Gelelectrophorietic analysis of phenol- extractable leaf portions from different viroid/host combinations. Arch. of Virol. 74:167-180.

Civan, P., and Svec, M. 2009. Genome-wide analysis of rice (Oryza sativa L. subsp. japonica) TATA box and Y Patch promoter elements. Genome/National Research Council Canada.52:294–297.

Clark, S. M., Vaitheeswaran, V., Ambrose, S. J., Purves, R. W., and Page, J. E. 2013. Transcriptome analysis of bitter acid biosynthesis and precursor pathways in hop (Humulus lupulus). BMC Plant Biol. 13:12.

Collard, C. Y., and Mackill, D. J. 2008. Marker-assisted selection: an approach for precision plant breeding in the twenty-first century. Phil. Trans. R. Soc. 363:557–572.

Conejero, V., Picazo, I., and Segado, P. 1979. Citrus exocortis viroid (CEV): protein alterations in different hosts following viroid infection. Virology 97:454-456.

Cuperus, J. T., Fahlgren, N., and, Carrington J. C. 2011. Evolution and functional diversification of MIRNAgenes. Plant Cell. 23:431–42.

Datta, K., Velazhahan, R., Oliva, N., Ona, I., Mew, T. 1999. Over-expression of the cloned rice thaumatin-like protein (PR-5) gene in transgenic rice plants enhances environmental friendly resistance to Rhizoctonia solani causing sheath blight disease. Theor Appl Genet 98: 1138–1145.

120

Deihimi, T., Niazi, A., Ebrahimi, M., Kajbaf, K., Fanaee, S., Bakhtiarizadeh, M. R., and Ebrahimie, E. 2012. Finding the undiscovered roles of genes: an approach using mutual ranking of coexpressed genes and promoter architecture-case study: dual roles of thaumatin like proteins in biotic and abiotic stresses. Springer Plus, 1, 30. http://doi.org/10.1186/2193- 1801-1-30.

De Keukeleire, J., Ooms, G., Heyerick, A., Roldan-Ruiz, I., Van Bockstaele, E., and De Keukeleire, D. 2003. Formation and accumulation of alpha-acids, beta-acids, desmethylxanthohumol, and xanthohumol during flowering of hops (Humulus lupulus). J. Agric. Food Chem. 16;51(15):4436-41.

Deng, W., and Roberts, S. G. 2005. A core promoter element downstream of the TATA box that is recognized by TFIIB. Genes and development 19:2418–2423.

Diebold, R., Schuster, J., Däschner, K., and Binder, S. 2002. The branched-chain amino acid transaminase gene family in Arabidopsis encodes plastid and mitochondrial proteins. Plant Physiol. 129:540–550.

Dhekney, S. A., Li, Z. T., Gray, D. J. 2010. Genetically engineered grapevines expressing a cisgenic Vitis vinifera thaumatin-like protein exhibit fungal resistance and improved post-harvest characteristics. 12th World congress of the International Association for Plant Biotechnology, St. Louis, June 6th-11th2010.

Diener, T. O. 1971. Potato spindle tuber “virus” IV. A replicating, low molecular weight RNA. Virology 45:411-428.

Diener, T. O. 1974. Viroids: the smallest known agents of infectious disease, Annu. Rev. Microbiol. 28:23–40.

Diener, T. O. 2003. Discovering viroids, a personal perspective. Nature Reviews Microbiology 1, 75-80.

Domingo, E., Escarmis, C., Sevilla, N., Moya, A., Elena, S. F., Quer, J., Novella S. I., and Holland, J. J. 1996. Basic concepts in RNA virus evolution, FASEB J. 10:859–864.

Eamens, A. L., Smith, N. A., Dennis, E. S., Wassenegger, M., Wang, M. B., 2014. In Nicotiana species, an artificial microRNA corresponding to the virulence modulating region of potato spindle tuber viroid directs RNA silencing of a soluble inorganic pyrophosphatase gene and the development of abnormal phenotypes. Virology 450–451, 266–277.

121

Eastwell, K. C., and Nelson, M. E. 2007. Occurrence of viroids in commercial hop (Humulus lupulus L.) production areas of Washington State. Online. Plant Health Progress. doi:10.1094/PHP-2007-1127-01-RS.

Edwardson, J. R. 1952. Hops: Their Botany, History, Production and Utilization. Economic Botany. 6: 160-175.

Elena, S. F., Dopazo, J., Flores, R., Diener, T. O., and Moya, A. 1991. Phylogeny of viroids, viroid like satellite RNAs, and the viroid like domain of hepatitis delta virus RNA. Proc. Natl. Acad. Sci. U.S.A. 88(13):5631–4.

Emanuelsson, O., Nielsen, H., Brunak, S., and von Heijne, G. 2000. Predicting subcellular localization of proteins based on their N-terminal amino acid sequence. J. Mol. Biol. 300:1005–1016.

Epple, P., Apel, K., and Bohlmann, H. 1995. An Arabidopsis thaliana thionin gene is inducible via a signal transduction pathway different from that for pathogenesis-related proteins. Plant. Physiol. 109:813-820.

Eulalio, A., Huntzinger, E., and Izaurralde, E. 2008. Getting to the root of miRNA-mediated gene silencing. Cell 132:9–14.

Eulgem, T. 2006. Dissecting the WRKY Web of Plant Defense Regulators. PLoS Pathog. 2(11): e126. doi:10.1371/journal.ppat.0020126.

Fierens, E., Rombouts, S., Gebruers, K., Goesaert, H., Brijs, K., Beaugrand, J., Volckaert, G., Van Campenhout, S., Proost, P., Courtin, C. M., Delcour, J. A. 2007. TLXI, a novel type of xylanase inhibitor from wheat (Triticum aestivum) belonging to the thaumatin family. Biochem. J. 403:583-91.

Flores, R. P., Serra, S. Minoia, F., Di Serio, B., and Navarro, J. 2012. Viroids: from genotype to phenotype just relying on RNA sequence and structure motifs. Front. Microbiol. 3:217.

Flores, R., Delgado, S., Rodio, M. E., Ambrós, S. Hernández, C., and Serio, F. D. 2006. Peach latent mosaic viroid: not so latent, Mol. Plant Pathol. 7:209–221.

Flores, R., DiSerio, F., and Hernández, C. 1997. Viroids: the noncoding genomes. Semin. Virol. 8: 65–73.

Flores, R., Navarro, J. A., de la Pena, M., Navarro, B. S., and Ambrós, A. 1999. Viroids with hammerhead ribozymes: some unique structural and functional aspects with respect to other members of the group. Biol. Chem. 380:849–854.

122

Frey, M., Chomet, P., Glawischnig, E., Stettner, C., Grün, S., Winklmair, A., Eisenreich, W., Bacher, A., Meeley, R. B., Briggs, S. P., Simcox, K., and Gierl, A. 1997. Analysis of a Chemical Plant Defense Mechanism in Grasses. Science. 277:696-699.

Fujimori, S., Washio, T., and Tomita, M. 2005. GC-compositional strand bias around transcription start sites in plants and fungi. BMC Genomics. 6(1):26.

Garcia, D. 2008. A miRacle in plant development: Role of microRNAs in cell differentiation and patterning. Semin. Cell Dev. Biol. 19:586–595.

Garcia-Olmedo, F., Molina, A., Segura, A., and Moreno, M. 1995. The defensive role of nonspecific lipid-transfer proteins in plants. Trends Microbiol 3:72-74.

Georgensgmuend, H. M. 2013. Barth Report. Joh Barth, Sohn GmbH and Co KG. Nuremberg.

Gomez, G., Pallas, V., 2001. Identification of an in vitro ribonucleoprotein complex between a viroid RNA and a phloem protein from cucumber plants. Mol Plant–Microbe Interact. 14, 910–913.

Gomez, G., Martınez, G., and Pallas, V. 2008. Viroid-Induced Symptoms in Nicotiana benthamiana Plants Are Dependent on RDR6 Activity. Plant Physiology. 148:414–423.

Gozmanova, M., Denti, M. A., Minkov, I. N., Tsagris, M., Tabler, M., 2003. Characterization of the RNA motif responsible for the specific interaction of potato spindle tuber viroid RNA (PSTVd) and the tomato protein Virp1. Nucleic Acids Res. 31, 5534–5543.

Hammond-Kosack, K. E., and Jones J. G. 1996. Resistance gene-dependent plant defense responses. The Plant Cell 8:1773-1791.

Hancock, D., and Holler, S. 1993. Strategic laboratory sampling-part 12. Popul. Med News 6:1-8.

Haunold, A. 1971. Cytology, sex expression, and growth of a tetraploid X diploid cross in hop (Humulus lupulus L.). Crop Sci. 11:868-871.

Haunold, A. 1972. Self fertilization in a normally dioecious species, Humulus lupulus L. J. of Hered. 63:283-286.

Havecker, E. R., Wallbridge, L. M., Hardcastle, T. J., Bush, M. S., and Kelly, K. A. 2010. The Arabidopsis RNA directed DNA methylation Argonautes functionally diverge based on their expression and interaction with target loci. Plant Cell 22:321–34.

123

Herr, A. J., Molnar, A., Jones, A., and Baulcombe, D. C. 2006. Defective RNA processing enhances RNA silencing and influences flowering of Arabidopsis. Proc Natl Acad Sci USA 103:14994–5001.

Higuchi, N., Ito, Y., Kato, J., Ogihara J, and Kasumi, T. 2015. NP24 induces apoptosis dependent on caspase-like activity in Saccharomyces cerevisiae. J. Biosci Bioeng [Epub ahead of print Nov. 15, 2015. Howard, G. A. 1964. The constituents and brewing behavior of hops. Hops World Crops Books, A. H. Burgess, ed., Interscience, New York and Leonard Hill, London. pp. 255-269.

Hsu, J. Y., Juven-Gershon, T., Marr, M. T., Wright, K. J., Tjian, R., and Kadonaga, J. T. 2008. TBP, Mot1, and NC2 establish a regulatory circuit that controls DPE-dependent versus TATA-dependent transcription. Genes Dev. 22:2353–2358.

Hutchins, C. J., Rathjen, P.D., Forster, A. C., and Symons, R. H. 1986. Self-cleavage of plus and minus RNA transcripts of avocado sunblotch viroid, Nucleic Acids Res. 14:3627–3640.

Jacob F. and Monod J. 1961. Genetic regulatory mechanisms in the synthesis of proteins J. Mol Biol. 3:318-56.

Jacob, F., Perrin, D., Sanchez, C., and Monod, J. 1960. Operon: a group of genes with the expression coordinated by an operator; C. R. Hebd Seances Acad Sci. 29:250:1727-9.

Joshi, C. P. 1987. An inspection of the domain between putative TATA box and translation start site in 79 plant genes. Nucleic Acids Res. 15:6643–6653.

Juven-Gershon, T., and Kadonaga, J. T. 2010. Regulation of gene expression via the core promoter and the basal transcriptional machinery. Developmental Biology 339:225–229.

Katiyar-Agarwal, S., Morgan, R., Dahlbeck, D., Borsani, O., Villegas, Jr., A., Jian-Kang, Z., Staskawicz, B., Jin, j., and Hailing. 2006. A pathogen-inducible endogenous siRNA in plant immunity. P. Natl. Acad. Sci. USA. 103(47):18002-18007.

Kawaguchi-Ito, Y., Li, S., Tagawa, M., Araki, H., Goshono, M., Yamamoto, S., Tanaka, M., Narita, M.,Tanaka, K., Liu, S., Shikata, E., and Sano, T. 2009. Cultivated Grapevines Represent a Symptomless Reservoir for the Transmission of Hop Stunt Viroid to Hop Crops: 15 Years of Evolutionary Analysis. PLoS ONE 4(12): e8386. doi:10.1371/journal.pone.0008386.

Keese, P., and Symons, R. H. 1985. Domains in viroids: evidence of intermolecular RNA rearrangements and their contribution to viroid evolution. Proc. Natl. Acad. Sci. U.S.A. 82:4582–4586.

124

Kochevenko, A., Klee, H. J., Fernie, A. R., and, Araújo, W. L. 2012. Molecular identification of a further branched-chain aminotransferase 7 (BCAT7) in tomato plants. J. Plant Physiol. 169:437–443.

Kojima M., Murai, M., and Shikata, E. 1983. Cytopathic Changes In Viroid-Infected Leaf Tissues. J. Fac. Agric. Hokkaido Univ. 61:219.

Komarnytsky, S., and Borisjuk, N. 2003. Functional Analysis of Promoter Elements in Plants. Genetic Engineering. 25:113-141.

Kovalskaya, N., and Hammond, R.W., 2014. Molecular biology of viroid–host interactions and disease control strategies. Plant Sci. 228:48–60.

Kumari, S., and Ware, D. 2013. Genome-Wide Computational Prediction and Analysis of Core Promoter Elements across Plant Mononcots and Dicots PLOS One. DOI: 10.1371/journal.pone.0079011.

Kusano, N., and Shimomura, K. 1997. Selection of PCR primers and a simple extraction method for detection of hop stunt viroid-plum in plum by reverse transcription-polymerase chain reaction. Annals of the Phytopath. Soc. of Japan 63:119-123.

Lagrimini, L. M., Burkhart, W., Moyer, M., and Rothstein, S. 1987. Molecular cloning of complementary DNA encoding the lignin-forming peroxidase from tobacco: Molecular analysis and tissue-specific expression. Proc. Natl. Acad. Sci. USA 84:7542-7546.

Liu, J-J, Sturrock, R., and Ekarmoddoullah, A. K. M. (2010). The superfamily of thaumatin-like proteins: its origin, evolution, and expression towards biological function. Plant Cell Reports 29:419-436.

Liu, J-J, and Ekarmoddoullah, A. K. M. 2006. The family 10 of plant pathogenesis-related proteins: Their structure, regulation, and function in response to biotic and abiotic stresses. Physiological and Molecular Plant Pathology 68:3–13.

Liu, D., Raghothama, K. G., Hasegawa, P. M., and Bressan, R. A. 1994. Osmotin overexpression in potato delays development of disease symptoms. Proc. Natl. Acad. Sci. U.S.A. 91: 1888–1892.

Mackintosh, C. A., Lewis, J., Radmer, L. E., Shin, S., Heinen, S. J. 2007. Overexpression of defense response genes in transgenic wheat enhances resistance to Fusarium head blight. Plant Cell Rep. 26:479–488.

125

Makino, S., Chang, M. F., and Shieh, C. K. 1987. Molecular cloning and sequencing of a human hepatitis delta (delta) virus RNA. Nature 329:343–6.

Maloney, G. S., Kochevenko, A., Tieman, D., Tohge, T., Krieger, U., Zamir, D., Taylor, M .G., Fernie A. R., and Klee, J. 2010. Characterization of the branched-chain amino acid aminotransferase enzyme family in tomato. Plant Physiol. 153:925–936.

Maniataki, E., Tabler, M., and Tsagris, M. 2003. Viroid RNA systemic spread may depend on the interaction of a 71-nucleotide bulged hairpin with the host proteinVirP1. RNA. 9:346– 354.

Martin, W.H. 1922. “Spindle tuber”, a new potato trouble. Hints to potato growers, N. J. State Potato Assoc. 3:8.

Martinez, G., Castellano, M., Tortosa, M., Pallás, V., and Gómez, G., 2014. A pathogenic non- coding RNA induces changes in dynamic DNA methylation of ribosomal RNA genes in host plants. Nucleic Acids Res. 42:1553–1562.

Matzke, M., Kanno, T., Daxinger, L., Huettel, B., and Matzke, A. J. 2009. RNA-mediated chromatin-based silencing in plants. Curr. Opin. Cell Biol. 21:367–76.

McInnes, J. L., and Symons, R. H. 1991. Comparative structure of viroids and their rapid detection using radioactive and nonradioactive nucleic acid probes. Viroids and Satellites: Molecular Parasites at the Frontier of Life, ed. K. Maramorosch Boca Raton, FL, CRC Press pp. 21–58.

Meilgaard, M. 1960. Hop analysis, cohumulone factor and the bitterness of beer: review and critical evaluation. J. Inst. Brew. 66:35-50.

Metraux, J. P., Burkhart, W., Moyer, M., Dincher, S., Middlesteadt, W., Williams, S., Payne, G., Carnes, M., and Ryals, J. 1989. Isolation of a complementary DNA encoding a chitinase with structural homology to a bifunctional lysozyme/chitinase. Proc. Natl. Acad. Sci. U.S.A. 86:896-900.

Molina, C., and Grotewold, E. 2005. Genome wide analysis of Arabidopsis core promoters BMC Genomics. 6:25.

Momma, T., and Takahashi, T. 1982. Ultrastructure of Hop Stunt Viroid-Infected Leaf Tissue. Phytopathology 104: 211-221.

126

Mühlbach, H. P., and Sänger, H. L. 1979. Viroid replication is inhibited by alpha-amanitin. Nature. 278:185-8.

Murakami, K., Elmlund, H., Kalisman, N., Bushnell, D. A., Adams, C. M., Azubel, M., Elmlund, D., Levi-Kalisman, Y., Liu, X., Gibbons, B. J., Levitt, M., and Kornberg, R. D. 2013. Architecture of an RNA Polymerase II Pre-Initiation Complex. Science 342:6159.

Muthamilarasan, M., and Prasad, M. 2013. Plant innate immunity: an updated insight into defense mechanism. J. Biosci. 2013 38(2): 433-49.

Nagel, J., Culley, L., Lu, Y., Liu, E., Matthews, P. D., Stevens, J. F., and Page, J. E. 2008. EST analysis of hop glandular trichomes identifies an O-methyltransferase that catalyzes the biosynthesis of xanthohumol. Plant Cell 20:186–200.

National Agricultural Statistics Service (NASS). 2012, 2011. Statistical Report Agriculture Status Board, USDA.

Natsume, S., Takagi, H., Shiraishi, A., Murata, J., Toyonaga, H., Patzak, J., Takagi, M., Yaegashi, H., Uemura, A., Yoshida, K., Krofta, K., Satake, H., Terauchi, R., and Ono, E. 2015. The Draft Genome of Hop (Humulus lupulus), an Essence for Brewing. Plant Cell Physiol. 56:428-441.

Narasimhan, M.L., Coca, M.A., Jin, J., Yamauchi, T., Ito, Y., Kadowaki, T., Kim, K.K., Pardo, J.M., Damsz, B., Hasegawa, P.M., Yun, D.J., Bressan, R.A. (2005). Osmotin is a homolog of mammalian adiponectin and controls apoptosis in yeast through a homolog of mammalian adiponectin receptor. Mol Cell 17(2):171-80.

Navarro, B., Gisel, A., Rodio, M. E., Delgado, S., Flores, R., and Di Serio, F., 2012. Small RNAs containing the pathogenic determinant of a chloroplast-replicating viroid guide the degradation of a host mRNA as predicted by RNA silencing. Plant J. 70:991–1003.

Neve, R.A. 1986. Centenary Review Hop Breeding Worldwide—its Aims and Achievements. J. Inst. Brew. 92:21-24.

Neve, R. A. 1991. Hops. Chapman and Hall. London.

Nickerson, G. A., Williams, P. A., and Haunold, A. 1986. Varietal differences in the proportions of cohumulone, adhumulone, and humulone in hops. J. Am. Soc. Brew. Chem. 44:91- 94.

127

Niemeyer, H. M. 1988. Hydroxamic acids (4-hydroxy-1,4-benzoxazin-3-ones) defense chemicals in the Gramineae. Phytochemistry. 27:3349-3358.

Ocamb, C. M. 2016. Hop (Humulus lupulus)-Fusarium Canker Wilt. Pacific Northwest Plant Disease Management Handbook. pnwhandbooks.org/plantdisease/node/3363 Pscheidt, J.W., and Ocamb, C.M. (Senior Eds.). 2016.

Ohno, T., Takamatsu, N., Meshi, T., and Okada, Y. 1983. Hop stunt viroid: molecular cloning and nucleotide sequence of the complete cDNA copy. Nucleic Acids Res. 11: 6185−6197.

Ohto, M., Onai, K., Furukawa, Y., Aoki, E., Araki, T., and Nakamura, K. 2001. Effects of sugar on vegetative development and floral transition in Arabidopsis. Plant Physiol. 127:252– 261.

Ohto, M. A., Fischer, R. L., Goldberg, R. B., Nakamura, K., and Harada, J. J. 2005. Control of seed mass by APETALA2. Proc. Natl. Acad. Sci. U.S.A. 102:3123–3128.

Okada, Y., and Ito, K. 2001. Cloning and analysis of valerophenone synthase gene expressed specifically in lupulin gland of hop (Humulus lupulus L.). Biosci. Biotechnol. Biochem. 65(1):150-5. Ong, C. T., and Corces, V. G. 2011. Enhancer function: new insights into the regulation of tissue-specific gene expression. Nature Rev. Genet. 12:283–293.

Osmond, R. I., Hrmova, M., Fontaine, F., Imberty, A., and Fincher, G. B. 2001. Binding interactions between barley thaumatin-like proteins and (1,3)-beta-D-glucans. Kinetics, specificity, structural analysis and biological implications. Eur. J. Biochem. 268:4190- 4199.

Owens, R. A., Blackburn, M., and Ding, B. 2001. Possible involvement of the phloem lectinin long-distance viroid movement. Mol. Plant Microbe Interact. 14:905–909.

Owens, R. A., Tech, K. B., Shao, J. Y., Sano, T., and Baker, C. J. 2012. Global analysis of tomato gene expression during Potato spindle tuber viroid infection reveals a complex array of changes affecting hormone signaling. Mol. Plant Microbe. Interact. 25:582–598.

Paniego, N. B., Zuurbier, K. W. M., Fung, S. Y., van der Heijden, R., Scheffer, J. J. C., and Verpoorte, R. 1999. Phlorisovalerophenone synthase, a novel polyketide synthase from hop (Humulus lupulus L.) cones. Eur. J. Biochem. 262:612–616.

Pardee, A. B., Jacob, F., and Monod, J. 1959. The genetic control and cytoplasmic expression of "inducibility" in the synthesis of B-galactosidase by E. coli. J. Mol. Biol. 1:165-178.

128

Park, C. J., An, J. M., Shin, Y. C., Kim, K. J., Lee, B. J., and Paek, K. H. 2004. Molecular characterization of pepper germin-like protein as the novel PR-16 family of pathogenesis related proteins isolated during the resistance response to viral and bacterial infection. Planta 219:797-806.

Patzak, J. 2005. Utilization of Molecular Biology Methods in Hop Breeding and Management in the Czech Republic. Acta. Hort. 668:67-74.

Pennacchio, L. A., Bickmore, W., Dean, A., Noberga, M. A., and Bejerano, G. 2013. Enhancers: five essential questions. Nature Reviews Genetics 14 (4):288-95.

Petre, B., Major, I., Rouhier, N., and Duplessis, S. 2011. Genome-wide analysis of eukaryote thaumatin-like proteins (TLPs) with an emphasis on poplar. BMC Plant Biology 11:33.

Qi, X., Bakht, S., Leggett, M., Maxwell, C., Melton, R., and Osbourn, A. 2004. A gene cluster for secondary metabolism in oat: implications for the evolution of metabolic diversity in plants. Proc. Natl. Acad. Sci. U.S.A. 101:8233–8238.

Qi, Y., and Ding, B. 2003. Inhibition of cell growth and shoot development by a specific nucleotide sequence in a noncoding viroid RNA. Plant Cell 15:1360–1374.

Riesner, D., Henco, K., Rokohl, U., Klotz, G., Kleinschmidt, A. K., Domdey, H., Jank, P., Gross, H. J., and Sänger, H. L. 1979. Structure and structure formation of viroids. J. Mol. Biol. 133:85–115.

Rodrigo, I., Vera, P., Tornero, P., Hernandez-Yago, J., and Conejero, V. 1993. cDNA Cloning of Viroid-lnduced Tomato Pathogenesis-Related Protein P23 Characterization as a Vacuolar Antifungal Factor. Plant Physiol. 102: 939-945.

Ruiz, F. A., Blanco-Portales, R., Mun-Blanco, R., and Caballero, J. 2011. The Strawberry Plant Defense Mechanism: A Molecular Review. Plant Cell Physiol. 52(11):1873–1903.

Ruiz-Ferrer, V., and Voinnet, O. 2009. Roles of Plant Small RNAs in Biotic Stress Responses. Annu. Rev. Plant Biol. 60:485–510.

Ryan, C. A. 1990. Protease inhibitors in plants: Genes for improving defenses against insects and pathogens. Annu. Rev. Phytopathol. 28:425-449.

Salgado, H., Moreno-Hagelsieb, G., Smith, T. F., and Collado-Vides. 2000. Operons in Escherichia coli: genomic analyses and predictions. Proc. Natl. Acad. Sci. U. S. A. 97(12):6652-6657.

129

Sano, O., Hataya, U., Shikata, E., Chou, M., and Okada. 1986. A Viroid Resembling Hop Stunt Viroid in Grapevines from Europe, the United States and Japan J. Gen.Virol.67:1673- 1678.

Sano, T., Ohshima, K., Uyeda, L., Shikata, E., Meshi, T., and Okada, Y. 1985. Nucleotide sequence of grapevine viroid: a grapevine isolate of hop stunt viroid. Proc. Japan Acad. 61(Ser .B) 265-268.

Sano, T., Uyeda, I., Shikata, E., Ohno, T., and Okada, Y. 1984. Nucleotide sequence of cucumber pale fruit viroid: homology to hop stunt viroid. Nucleic Acids Res. 12(8):3427–3434.

Schmidt, R., and Bancroft, I. 2011. Genetics and Genomics of the Brassicaceae. New York, NY. Springer Science and Business Media. 3:72.

Seefelder, S., Ehrmaier H., Schweizer, G., and Seigner, E. 2000. Male and female genetic linkage map of hops, Humulus lupulus. Plant Breeding 119: 249-255.

Shimura, K., Okada, A., Okada, K., Jikumaru, Y., Ko, K.W., Toyomasu, T., Sassa, T., Hasegawa, M., Kodama, O., Shibuya, N., Koga, J., Nojiri, H., and Yamane, H. 2007. Identification of a biosynthetic gene cluster in rice for momilactones. J. Biol. Chem. 282:34013–34018.

Shimura, H., Pantaleo, V., Ishihara, T., Myojo, N., Inaba, J., Sueda, K., Burgyán, J., and Masuta, C. 2011. A viral satellite RNA induces yellow symptoms on tobacco by targeting a gene involved in chlorophyll biosynthesis using the RNA silencing machinery. PLoS Pathog. 7:e1002021. Published online.

Singh, B. K., and Shaner, D. L. 1995. Biosynthesis of branched chain amino acids: from test tube to field. Plant Cell 7:935–944.

Singh, R. P., 1971. A local lesion host for potato spindle tuber virus. Phytopathology 61:1034–1035.

Small, E. 1978. A numerical and nomenclatural analysis of morphogeographic taxa of Humulus. Syst. Bot. 3:37–76.

Small, E. 1980. The relationships of hop cultivars and wild variants of Humulus lupulus. Can. J. Bot. 58:676–686.

Smith, N. A., Singh, S. P., Wang, M. B., Stoutjesdijk, P. A., Green, A. G., and Waterhouse, P. M. 2000. Gene expression: total silencing by intron-spliced hairpin RNAs. Nature 407: 319– 320.

130

Smith, N. A., Eamens, A. L., and Wang, M. B. 2011. Viral small interfering RNAs target host genes to mediate disease symptoms in plants. PLoS Pathog. 7:e1002022. Published online.

Spoel, S. H., and Dong, X. 2012. How do plants achieve immunity? Defense without specialized immune cells. Nature Reviews. 12:89-100.

Stajner, N., Cregeen, S., and Javornik, B. 2013. Evaluation of Reference Genes for RT-qPCR Expression Studies in Hop (Humulus lupulus L.) during Infection with Vascular Pathogen Verticillium albo-atrum. PLoS ONE 8(7):e68228.

Steenackers, B., De Cooman, L., and De Vos, D. 2015. Chemical transformations of characteristic hop secondary metabolites in relation to beer properties and the brewing process: A review Food Chemistry 172:742–756.

Stevens, R. 1967. The chemistry of hop constituents. Chem. Rev. 67:19-71.

Sue, M., Nakamura, C., and Nomura, T. 2011. Dispersed Benzoxazinone gene cluster: Molecular Characterization and Chromasomal Location. Plant physiology 157(3):985- 997.

Symons, R. H. 1981. Avocado sunblotch viroid: primary sequence and proposed secondary structure, Nucleic Acids Res. 9:6527–6537.

Terras, F. G., Schoofs, H., De Bolle, M. C., van Leuven F., Rees, S. B., Vanderleyden, J., Cammue B. A., and Broekaert, W.F. 1992 Analysis of two novel classes of plant antifungal proteins from radish (Raphanus sativus L.) seeds. J. Biol.Chem. 267:15301-15309.

Unger, R. W. 2004. Beer in the Middle Ages and the Renaissance. University of Pennsylvania Press. Pages 15-36.

Van Loon, L. C. 1997. Induced resistance in plants and the role of pathogenesis-related proteins. Eur. J. of Plant Path. 103:753-765.

Van Loon, L. C., Gerritsen, Y. M., and Ritter C. E. 1987. Identification, purification, and characterization of pathogenesis-related proteins from virus-infected Samsun NN tobacco leaves. Plant Mol. Biol. 9:593-609.

Van Loon, L. C., Rep, M., and Pieterse, C. M. 2006. Significance of inducible defense-related proteins in infected plants. Annu. Rev. Phytopathol. 44:135-62.

131

Van Loon, L. C., and Van Kammen, A. 1970. Polyacrylamide disc electrophoresis of the soluble leaf proteins from Nicotiana tabacum var ‘Samsun’ and Samsun NN. II. Changes in protein constitution after infection with tobacco mosaic virus. Virology. 40:199-211.

Van Loon, L. C., and Van Strien, E. A. 1999. The families of pathogenesis-related proteins, their activities, and comparative analysis of PR-1 type proteins. Phys. and Mol. Plant Path. 55:85-97.

Verzele, M. 1986. Centenary review: 100 years of hop chemistry and its relevance to brewing. J. Inst. Brew. 92:32-48.

Verzele, M., and De Keukeleire, D. 1991. Chemistry and analysis of hop and beer bitter acids. Elsevier, Amsterdam.

Voinnet, O. 2009. Origin, Biogenesis, and Activity of Plant MicroRNAs. Cell. 136(4):669 – 687.

Wang, G., and Dixon, R. A. 2009. Heterodimeric geranyl (geranyl) diphosphate synthase from hop (Humulus lupulus) and the evolution of monoterpene biosynthesis. Proc. Natl. Acad. Sci. 106:9914–9919.

Wang, G., Tian, L., Aziz, N., Broun, P., Dai, X., He, J., King, A., Zhao, P.X., and Dixon, R. A. 2008. Terpene biosynthesis in glandular trichomes of hop. Plant Physiol. 148:1254–1266.

Wang, Y., Wu, J., and Ding, B., 2014. A deeply conserved transcription factor regulates the replication of potato spindle tuber viroid RNA genome. In: XVIth Internatl. Congress of Virology, Montreal.

Wu, L., Zhou, H., Zhang, Q., Zhang, J., and Ni, F. 2010. DNA methylation mediated by a microRNA pathway. Mol. Cell. 38:465–75.

Wroblewski, T., Matvienko, M., Piskurewicz, U., Xu, H., Martineau, B., Wong, J., Govindarajulu, M. Kozik, A., and Michelmore, R. 2014. Distinctive profiles of small RNA couple inverted repeat-induced post-transcriptional gene silencing with endogenous RNA silencing pathways in Arabidopsis. RNA 20:1987–1999.

Yaguchi, S., and Takahashi, T. 1984. Response of Cucumber Cultivars and Other Cucurbitaceous Species to Infection by Hop Stunt Viroid. J. of Phytopathol. 109:21-31.

Yamamoto Y. Y., Yoshitsugu, T., Sakurai, T., Seki, M., Shinozak, K., and Obokata, J. 2009. Heterogeneity of Arabidopsis core promoters revealed by high-density TSS analysis. The Plant Journal. 60:350–362.

132

Yang, C., Bolotin, E., Jiang, T., Sladek, F.M., and Martinez, E. 2007. Prevalence of the initiator over the TATA box in human and yeast genes and identification of DNA motifs enriched in human TATA-less core promoters. Gene 389:52–65.

Yang, M. Q., van Velzen, R., Bakker, F. T., Sattarian, A., Li, D. Z., and Yi, T. S. 2013. Molecular phylogenetics and character evolution of Cannabaceae. Taxon. 62:473–485.

Yant, L., Mathieu, J., Dinh, T. T., Ott, F., Lanz, C., Wollmann, H., Chen, X., and Schmida, M. 2010. Orchestration of the floral transition and floral development in Arabidopsis by the bifunctional transcription factor APETALA2. The Plant Cell. 22: 2156–2170.

Yoshiharu, Y., Yamamoto, H. I., Minami, M., Junichi, O., Tetsuya, S., Masakazu, S., Motoaki S., Kazuo, S., and Tomoko, A. 2007. Identification of plant promoter constituents by analysis of local distribution of short sequences BMC Genomics. 8:67.

Zhou, X. J., Lu, S., Xu, Y. H., Wang, J. W., and Chen, X, Y. 2002. A cotton cDNA (GaPR-10) encoding a pathogenesis-related 10 protein with in vitro ribonuclease activity. Plant Sci. 162:629-636.

Zuurbier, K. W. M., Fung S. Y., Scheffer J. J. C., and Verpoorte R. 1995. Formation of aromatic intermediates in the biosynthesis of bitter acids in Humulus Lupulus. II Phytochemistry. 38:77-82.

133