DETECTION OF ERYSIPHE NECATOR (UNCINULA NECATOR)

WITH POLYMERASE CHAIN REACTION AND

SPECIES-SPECIFIC PRIMERS

By

JENNIFER SUSAN FALACY

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

MASTER OF SCIENCE IN PLANT PATHOLOGY

WASHINGTON STATE UNIVERSITY Department of Plant Pathology

DECEMBER 2003

ii

To the Faculty of Washington State University:

The members of the Committee appointed to examine the thesis of JENNIFER SUSAN FALACY find it satisfactory and recommend that it be accepted.

___________________________________ Chair

___________________________________

___________________________________

___________________________________

___________________________________

iii

ACKNOWLEDGMENTS

I would like to extend my appreciation to all whose contributions made this thesis

possible. First and foremost, I thank my advisor, Gary Grove, for his generosity and

amazing empowering attitude, enthusiasm, and humor which is contagious and

promotes laughter, levity, and teamwork in the lab. I am grateful to all my committee

members who guided and supported me through this research. A special thanks to

Dean Glawe who graciously donated a great deal of his time to teach me the

taxonomy of the powdery mildews and an appreciation of nomenclature. I would

have been lost without Richard Larsen and George Vandemark, who were an

endless source of molecular advice, trouble-shooting guidance, and encouragement.

I appreciate the assistance provided by Heather Galloway and Jeff Lunden in our lab,

who made light of even the worst day. The wonderful and helpful personalities of the

people in the lab and the entire experiment station made it a pleasure to work there.

I appreciate Terri Hughes for always being there; listening to me laugh and cry,

complain and rejoice, and deliberate decisions to death. Sue Ellen’s class would

have been impossible without you as a study partner. You saw me though some of

the greatest and worst days of this rite of passage. This time in my life will be

remembered as magical because you were there to share it with.

A special thanks to Duane Moser who introduced me to mycology, inspired me

toward this field and encouraged me throughout this process. I appreciate your

iv

patience, kind words and gentle hugs. Your editorial skills, time spent listening to me

practice seminars and providing insight contributed greatly to my confidence at the

thesis defense.

I am especially grateful to my family for patiently understanding all the time I

sacrificed spending with them in order to pursue this endeavor. In particular, I want

to thank Geri, Callie, and Mom for proofreading my assignments and making me

laugh with their creative comments.

v

DETECTION OF ERYSIPHE NECATOR (UNCINULA NECATOR)

WITH POLYMERASE CHAIN REACTION AND

SPECIES-SPECIFIC PRIMERS

Abstract

by Jennifer Susan Falacy, M.S. Washington State University

December 2003

Chair: Gary G. Grove A polymerase chain reaction (PCR) assay employing species-specific primers was

developed to differentiate Erysiphe necator (Uncinula necator) from other powdery

mildews common in the northwest United States. This assay is intended to be used

in conjunction with high efficiency air samplers for the addition of an inoculum

component to current grapevine powdery mildew risk assessment models. DNA was

extracted from mycelia, conidia, and/or cleistothecia that were collected from grape

leaves using a Burkard cyclone surface sampler. To differentiate E. necator from

closely related powdery mildew fungi, primer pairs Uncin144 and Uncin511 were

developed by aligning internal transcribed spacer (ITS) sequences from E. necator

and other powdery mildews and choosing regions unique to E. necator. The primers

generated amplicons specific to E. necator, but did not generate amplicons when

tested with powdery mildew species collected from 46 disparate hosts in 26 vascular

plant families. As a result of these tests, the amplification of a single 367 base pair

(bp) fragment using the primer pairs, and visualized by gel electrophoresis, was

vi

considered evidence of the presence of E. necator. Amplification products were

cloned and sequenced to verify the specificity of E. necator primers. This PCR-

based test could enable the detection of E. necator in field samples within hours of

collection, and when air sampling and identification protocols perfected, is expected

to result in significant improvements to grapevine powdery mildew risk assessment

models.

vii

LIST OF TABLES

1. Powdery mildews collected, identified and tested using PCR with primers

Uncin144 and Uncin511··························································································· 18

2. List of E. necator isolates from diverse geographic origins yielding amplification

products when PCR was performed using primers Uncin144 and Uncin511 ··········· 20

viii

LIST OF FIGURES

.

1. Internal Transcribed Spacer Region, a portion of DNA located between the

large and small ribosomal subunits. Uncin144 and Uncin511 are located

between the universal primers ITS1 and ITS4··············································· 21

2. Agarose gels showing amplification products from polymerase chain reaction

of internal transcribed spacer regions of selected powdery mildews using

universal primers ITS1 and ITS4 and E. necator-specific primer pair··········· 22

3. Agarose gel showing amplification products from polymerase chain reaction of

E. necator conidia added directly to the master mix ······································ 23

ix

ATTRIBUTIONS

G. G. Grove – Project leader, advisor, epidemiological advice, funding source.

R. C. Larsen- Molecular science specialist.

G. J. Vandemark- Cloning training and advice.

D. A. Glawe- Taxonomist. Provided training on the identification of the powdery mildews.

H. Galloway – Associate in Research. Ensured experiment continuation while Jennifer was in

Pullman attending classes.

x

TABLE OF CONTENTS

Page ACKNOWLEDGEMENTS ·······················································································iii ABSTRACT ············································································································v LIST OF TABLES····································································································vii LIST OF FIGURES ·································································································viii ATTRIBUTIONS······································································································ix INTRODUCTION ····································································································3 MATERIALS AND METHODS ···············································································7 Primer design ·······························································································7 Isolate identification······················································································7 Isolate collection ··························································································8 Field spore collection ···················································································9 DNA extraction ····························································································10 PCR assay ··································································································11 PCR on untreated spores ············································································11 DNA cloning and sequencing ······································································12 RESULTS ··············································································································13 DISCUSSION ·········································································································14 LITERATURE CITED ·····························································································24

xi

Detection of Erysiphe necator (Uncinula necator) with Polymerase Chain Reaction

and Species-Specific Primers

Jennifer S. Falacy, Gary G. Grove, and H. Galloway Irrigated Agriculture Research and Extension Center, Washington State University, Prosser, WA Richard C. Larsen and George J. Vandemark Vegetable and Forage Crops Production, Agricultural Research Service, United States Department of Agriculture, Prosser, WA D.A. Glawe Puyallup Research and Extension Center, Washington State University, Puyallup, WA Manuscript to be submitted to Phytopathology

1

Detection of Erysiphe necator (Uncinula necator) with Polymerase Chain

Reaction and Species-Specific Primers

J.S. Falacy, G.G. Grove, R.C. Larsen, G.J. Vandemark, D.A. Glawe, and

H. Galloway

First, second and sixth authors: Dept. Plant Pathology, Washington State University-Irrigated

Agriculture Research and Extension Center (IAREC), Prosser, WA 99350-9687. Third and fourth

author: Vegetable and Forage Crops Production, Agricultural Research Service, United States

Department of Agriculture, Prosser, WA 99350-9687. Fifth author Dept. Plant Pathology, Washington

State University- Puyallup Research and Extension Center, Puyallup, WA 98371-4571.

A polymerase chain reaction (PCR) assay employing species-specific primers was

developed to differentiate Erysiphe necator (Uncinula necator) from other powdery

mildews common in the northwest United States. This assay is intended to be used

in conjunction with high efficiency air samplers for the addition of an inoculum

component to current grapevine powdery mildew risk assessment models. DNA was

extracted from mycelia, conidia, and/or cleistothecia that were collected from grape

leaves using a Burkard cyclone surface sampler. To differentiate E. necator from

closely related powdery mildew fungi, primer pairs Uncin144 and Uncin511 were

developed by aligning internal transcribed spacer (ITS) sequences from E. necator

and other powdery mildews and choosing regions unique to E. necator. The primers

generated amplicons specific to E. necator, but did not generate amplicons when

2

tested with powdery mildew species collected from 46 disparate hosts in 26 vascular

plant families. As a result of these tests, the amplification of a single 367 base pair

(bp) fragment using the primer pairs Uncin144 and Uncin511, and visualized by gel

electrophoresis, was considered evidence of the presence of E. necator.

Amplification products were cloned and sequenced to verify the specificity of E.

necator primers. This PCR-based test could enable the detection of E. necator in

field samples within hours of collection, and when air sampling and identification

protocols perfected, is expected to result in significant improvements to grapevine

powdery mildew risk assessment models.

3

INTRODUCTION

Powdery mildew, caused by Erysiphe necator Schw. [Uncinula necator (Schw.) Burr.]

(Ascomycotina, Erysiphales), is the most economically significant disease of

viniferous grapes in the Pacific Northwest. The disease can negatively affect wine

pH, aroma, and flavor and can predispose berries to infection by other pathogens

such as Botrytis sp. (Gubler, 1996; Ough and Berg, 1979; Gadoury et al., 2001;

Pearson, 1988). Severe infestations can reduce vigor and winter hardiness (Pool et

al., 1984; Northover and Homeyer, 2001; Pearson, 1988; Grove, 2000). Attempts to

manage this disease have resulted in excessive chemical usage and labor costs

(Miazzi et al., 1997; Grove, 2003). In recent years fungicide usage has been slightly

reduced through the use of risk assessment models (Gubler, 1996). Existing risk

assessment models based on temperature and assumed inoculum presence and

activity are of limited value if the pathogen is not present in all active vineyards.

Limitations of these models can prevent growers from responding quickly to disease-

conducive meteorological conditions in order to prevent intensification of epidemics

(Jarvis et al., 2002). Fear of epidemic development based on assumed inoculum

presence and disregard of the significance of meteorological conditions can lead

growers to spray according to host phenology rather than in response to actual

disease-conducive conditions increasing risk. In some years fungicide applications

made according to the criteria provided by existing models may be much earlier than

necessary (Grove, 2003).

4

Accurate, timely detection of the pathogen’s presence could result in a more reliable

assessment of the risk of epidemic development. More dependable risk assessment

models would allow growers to utilize them and in doing so save time and expense

through reduced fungicide usage. This reduction would have economic and

environmental benefits. Detection of the pathogen through diagnosis of early

disease symptoms, such as a slight difference in varietal-dependant foliar texture or a

patchy lack of sheen on infected leaves, can be difficult to accomplish on varieties

with pubescent leaves or in poor lighting conditions. Such difficulties make it

impossible to employ effective powdery mildew management programs in a timely

manner based solely on disease scouting. The advanced symptoms of this disease,

including grayish-white powdery patches on both surfaces of leaves and

subsequently on berries, are easily recognized and diagnosed. However, the disease

is difficult to manage at this stage.

Conidia and ascospores of E. necator are dispersed primarily by air currents

(Hammett and Manners, 1974; Willocquet, et al., 1998). Although detection of

airborne spores of E. necator would be useful in establishing the presence of the

pathogen, traditional approaches of trapping and identifying powdery mildew fungi

are not practical for assessing the risk of epidemic development. Conventional

identification of Erysiphales relies upon microscopic assessment of morphological

characters (Braun, 1987; Braun 1995; Braun et al., 2002). Early detection would

require the identification of airborne spores that may not readily display the

morphological characters required for proper classification. This process is time

5

consuming, error prone, and requires the expertise of a person familiar with the

morphology of powdery mildews. The only isolation method successful in gaining

pure cultures of this obligate biotroph requires single-conidium transfers from a field

sample to surface sterilized detached leaves or leaf disks. Growing of a colony using

this technique takes up to two weeks to visualize evidence of colony growth with the

naked eye (Olmstead et al., 2000). Such approaches would not be practical for

monitoring large plantings for presence of the disease when rapid identification of the

pathogen would be required. However, the use of a device capable of sampling

large volumes of air, conidia, and ascospores used in conjunction with a sensitive

molecular-based diagnostic tool could allow for the quick and reliable detection of

pathogens early in the progress of an epidemic and growing season. The resulting

detection information could then be incorporated into existing forecasting models to

facilitate near real-time disease management decisions.

PCR-based diagnostic tools have been developed to identify fungi, bacteria, and

viruses for applications in food safety, medical, animal and crop sciences

(Venkateswaran et al., 1997; Williams et al., 2001; Kong et al., 2003; Rampersad and

Umaharan, 2003; Jimenez et al., 2000; Stark et al., 1998). The advantages of cost

savings, precision, and accuracy in identifying causal agents of disease have added

to the popularity of this useful technique (Kong et al., 2003; Levesque, 2001). Very

little research has focused on molecular-based detection of powdery mildew species,

and no studies have attempted the collection, detection, and identification of airborne

powdery mildew spores in an agricultural setting using PCR. However, the literature

6

contains a sizeable body of work characterizing powdery mildew fungi at the

molecular level that can provide a basis for such an effort (Takamatsu et al., 1998;

Hirata et al., 1996; Saenz and Taylor, 1999). Previous research has focused on the

Internal Transcribed Spacer (ITS) region of ribosomal DNA which is located between

the 18s and 28s subunit genes and repeated numerous times (Fig. 1.); (White et al.,

1990). Hirata (1996) concluded that while the rDNA ITS region is conserved, it is

sufficiently variable enough to facilitate phylogenetic studies of closely related

species. Several recent phylogenetic studies have led to sequencing of the ITS

region of several Erysiphaceous fungi (Takamatsu et al., 1998; Hirata et al., 1996;

Saenz and Taylor, 1999). ITS sequences from many of these organisms are

available in GenBank (Altschul et al., 1997) allowing for easy comparison of available

sequences. A further advantage of working with the ITS region is that several

hundred copies of this region exist per individual cell, making it easier to amplify the

region with PCR from small amounts of material (such as spores) than when using

non-repeated regions of the genome (Lee and Taylor, 1990). This region of the

genome was the focus of this study because of the lower detection threshold gained

as a result of targeting this repeated ribosomal sequence.

The objectives of the present study were to: (i) develop a PCR assay that would

consistently detect and distinguish E. necator from other powdery mildews occurring

in the Pacific Northwest, (ii) reliably detect E. necator from air samples made in a

field environment, (iii) evaluate the sensitivity of the assay.

7

MATERIALS AND METHODS

Primer design. The 45 powdery mildew sequences deposited with GenBank by

Saenz (1999) were aligned using Clustal W software (Thompson et al., 1994). PCR

primers were designed from conserved sequence fragments unique to E. necator.

Aligned areas unique to E. necator were selected as potential primer locations using

Primer Designer 4 (Sci-Ed, Cary, NC) software. Primers Uncin144 and Uncin511 are

nested between universal primers ITS1 and ITS4 (White et al., 1990) (Fig.1).

Isolate identification. Mycelia, conidia, and/or cleistothecia from powdery mildew

fungi were collected from infected leaves of native, introduced, horticultural, and

agricultural plants representing 46 different plant species within 26 families. Each

fungus was identified on the basis of host genus and fungal morphology.

Microscopic features used to distinguish species included: conidia size and shape

conidiophores; foot cells; appressoria; ascocarp (cleistothecia) size, shape, and

appendage type; number and shape of asci; and number and shape of ascospores.

Not all of these features were available for all of the fungi identified. The fungi were

identified using the taxonomic system of Braun (1987, 1995), with potential

discrepancies noted (Table 1). Braun and Takamatsu (Braun and Takamatsu, 2000;

Braun et al,. 2002) recently suggested changing genus concepts for Erysiphaceous

fungi. Table 1 lists names applied to fungi included in this study, giving both the

8

commonly used scientific names as well as those included in recent nomenclatural

proposals (Braun and Takamatsu 2000; Braun et al., 2002).

Isolate collection. Fungal material (conidia, cleistothecia, and/or mycelia) was

collected from host leaves using a Burkard cyclonic surface sampler (Burkhard Mfg.

Co., Rickmansworth Hertfordshire, Eng.) and deposited into 1.5 ml capless

microcentrifuge tubes with plug closures. DNA was extracted immediately from

collected material or the samples desiccated, flash frozen in liquid nitrogen, and

stored at –70°C as described previously (Stummer et al., 1999). Removable parts of

the cyclonic sampler were cleaned between sample collections by soaking in

Formula 409 (2-butoxyethanol)(Clorox, Pleasanton, CA) for 20 minutes and rinsing

with deionized water.

Samples originating from outside Washington State (Table 2) were preserved and

shipped in 70% or 95% ethanol. Tubes containing infected leaf material were

inverted several times to suspend conidia and other fungal material in the ethanol.

Leaf material was removed prior to samples being centrifuged at 1700 x g for 20 min

in a fixed-angle rotor using a clinical centrifuge (International Equipment Co.,

Needham Heights., MA) to concentrate the fungal material and spores. The

supernatant was discarded and the DNA extracted from the fungal pellet as

described below.

9

Field spore collection. In preliminary studies, two different spore traps were

evaluated to assess their efficacy in collecting fungal material in a vineyard to test

with the PCR detection technique. The devices were located 0.5 m downwind (from

prevailing winds) from a vineyard comprised of 3-year old Chardonnay and Riesling

grapes varieties located at WSU-IAREC, Prosser, WA. The vineyard was severely

infested with E. necator at the times of sampling.

Rotary-impaction studies. Two 5 cm by 1 mm glass rods coated in vacuum grease

(Dow Corning, Midland, MI) were secured in a battery-powered Rotorod® (Sampling

Technologies, Inc. Minnetonka, MN) spinning assembly. Air was sampled by

spinning the rods for 4-8 hours at approximately 2400 RPM. The glass rods were

then shattered into pieces small enough to fit in the microcentrifuge tubes, and DNA

extracted from collected spores utilizing the procedure described below.

High-efficiency cyclonic sampling. The Bioguardian® (Innovatek, Richland, WA)

samples 1000 liters of air per minute, sorts particles according to size, and deposits

particles 20 - 100 µm in a phosphate buffered saline buffer (136.9 mM NaCl, 1.47

mM KH2PO4, 8.04 mM Na2HPO4, 2.68mM KCL, 0.05% Triton 100 X). The

Bioguardian was programmed to collect for five minute periods at hourly intervals for

20-24 hours. Samples deposited in the collection buffer were centrifuged at 1700 x g

for 20 min in a fixed-angle rotor using a clinical centrifuge. Spores and other

particulate precipitate in the pellet were retained and the DNA extracted as described

below.

10

DNA extraction. DNA was extracted using a modification of the FastDNA (Bio 101,

Inc., Carlsbad, CA) protocol. Prior to the addition of the sample and the supplied

extraction buffer, 17 mg polyvinylpyrrolidone (PVP)(Sigma-Aldrich, St. Louis, MO)

was added to each microcentrifuge tube. Samples were homogenized for 30 s at an

intensity setting of 5.0 in the FastPrep FP 120 homogenizer (BIO 101/Savant, Vista,

CA). The process was repeated after a brief chilling period on ice. After

centrifugation for 10 min at 14,000 x g, 800 µl of the aqueous phase was transferred

to a 1.7 ml microcentrifuge tube and extracted with equal volume of phenol-

chloroform-isoamyl alcohol(1:1:24 v/v). The supernatant (600 µl) was transferred to

a clean 1.7 ml tube and the DNA bound to a matrix using the supplied binding buffer,

washed, and then eluted with 100 µl of sterile distilled water. DNA extracts were

stored at -20°C and diluted 1:6 with deionized water prior to amplification. The

protocol was followed for all powdery mildew samples with the exception of Medicago

sativa and Rubus ursinus powdery mildews. For the latter two species, the extraction

buffer consisted of an equal volume each of the supplied fungal and plant buffers

plus the addition of one-half volume each of the supplied PPS reagent and the PVP.

All DNA preparations were amplified using ITS1 and ITS4 universal primers

described by White et al., (1990), the results of which served as a positive control

indicating successful fungal DNA extraction.

11

PCR assay. PCR assays were conducted in 0.2 ml tubes consisting of 25 µl

reactions containing the PCR master mix: 20 mM Tris-HCl (pH 8.8), 10 mM KCL,

10mM (NH4)2SO4, 2mM MgSO4, 0.1% Triton X-100, 0.1mg/ml bovine serum albumin,

3 mM MgCl, 120 µM each of dATP, dTTP, dCTP, dGTP, 2.5 µM of each primer, 0.1

units of Pyrococcus furiosus (Pfu) DNA polymerase (Stratagene Corp., La Jolla, CA);

and 1 µl target DNA. A Biometra TGradient thermocycler (Whatman, Göttingen,

Germany) was used with a program of: 2 min initial denaturation at 96°C, followed

by 35 cycles of 30 s at 95°C, 30 s at 65°C (with universal primers) or 70°C (with

species-specific primers), 30 s at 70°C and a single final extension period of 7 min at

70°C. Amplified DNA was resolved on a 1% SeaKem® GTG® agarose gel

(BioWhittaker inc., Rockland, ME) in 1X Tris-borate-EDTA buffer (90mM Tris-borate

and 2mM EDTA). Amplification products were stained with ethidium bromide,

visualized under ultraviolet light, and recorded by digital image with an AlphaImager

2000 (Alpha Innotech, San Leandro, CA). The experiment was repeated at least

twice for each powdery mildew collected.

PCR on untreated spores. Short conidial chains were gathered from young fresh

infected grape leaves using a using a single eyelash attached to a glass Pasteur

pipette while viewing under a dissection microscope. Harvested conidia were

transferred from the leaf to the PCR master mix. The tubes containing the conidia

and master mix were spun briefly in a centrifuge followed by incubating at -20°C for

about 30 minutes. The PCR parameters were optimized to include a 6 min initial

denaturation at 96°C prior to the basic process described above. In addition the

12

number of PCR cycles was increased to 45. Freezing the master mix with the

spores, lengthening the initial denaturation step and increasing the number of PCR

cycles were performed with the intent of disrupting the spores.

DNA Cloning and Sequencing. PCR amplification products were excised from

agarose gels and purified using the GeneClean Turbo kit (Bio 101 Inc., Vista, CA).

Purified DNA fragments were ligated to the pCR®4-TOPO plasmid (TOPO TA;

Invitrogen Corp, Calsbad, CA) and transformed into DH5αT1 competent cells.

Selected colonies were incubated overnight in LB broth containing 10µg/ml

kanamycin in a shaking incubator at 37°C. Plasmid DNA was isolated from the

competent DH5αT1 cells by using the Wizard® Plus SV Miniprep DNA Purification

System (Promega, Madison, WI). Procedures for all preparations were conducted

according to the manufacturer’s instructions. Clones containing the PCR product

were identified by an additional PCR reaction performed on the purified plasmid DNA

using the primers Uncin144 and Uncin511. The DNA products were sequenced in

both directions using the deoxy-chain termination method by the Laboratory for

Biotechnology and Bioanalysis, School of Molecular Biosciences, Washington State

University.

13

RESULTS

Alignment of the complete ITS regions of the 45 powdery mildew species sequenced

by Saenz and Taylor revealed both highly conserved and low consensus variable

regions (Saenz and Taylor, 1999). Primers Uncin144 and Uncin511 were selected

because of their high specificity to E. necator.

Uncin144 (Forward) CCGCCAGAGACCTCATCCAA

Uncin511 (Reverse) TGGCTGATCACGAGCGTCAC

A NCBI-Blast2 (Altschul, 1997) search of our primer sequences showed that the

amplification product generated as a result of PCR with primers Uncin144 and

Uncin511 shared 100% homology with the E. necator (AF011325) sequence

deposited in Genbank. PCR using the universal primers, ITS1 and ITS4, yielded

amplification products between 500 -600 bp from all powdery mildews listed in Table

1 indicating that the DNA was of sufficient quality for amplification experiments. The

presence on agarose gels of the expected 367 bp PCR amplification product was

evidence of detection of E. necator when using primers Uncin144 and Uncin511.

The use of primers Uncin144 and Uncin511 resulted in amplification products from

only E. necator DNA but not from 35 species of powdery mildews (9 genera)

associated with the 46 host species representing 26 families of vascular plants other

than Vitis sp. (Table 1). Amplifications were successful from all E. necator isolates

collected regardless of geographic origin (Table 2). A test was considered successful

14

for differentiating E. necator from other Erysiphaceous fungi only if 1) amplification

with universal primers ITS1 And ITS4 yielded product for each powdery mildew DNA

sample listed in table 1, and 2) that PCR with primers Uncin144 and Uncin511

amplified only E. necator while failing to amplify the others (Fig. 2).

This protocol was also successful at identifying E. necator when conidia of the

pathogen were added directly to the PCR mix. In each of three experiments with

nine replicates E. necator conidia was detected with the following accuracy: five

conidia per reaction were detected in 89 % of the trials; two conidia were detected in

100 % of the trials; and one conidium was detected in 67% of the trials.

Both the Rotorod® and Bioguardian® air sampling devices provided efficient means

for collecting powdery mildew spores without interfering with subsequent DNA

extraction and PCR procedures. PCR products obtained from samples collected by

these devices revealed the presence of E. necator using primers Uncin144 and

Uncin511 by yielding amplification products of the expected size (Fig. 2C and D,

lane 7).

DISCUSSION

Results of this study indicated that primers Uncin144 and Uncin511 were specific for

E. necator regardless of its geographic origin. The PCR-based assay was able to

detect and differentiate (within hours of collection) E. necator DNA from that of other

15

Erysiphaceous DNA individually as well as in vineyard air samples containing a

background of unidentified airborne spores. This tool could facilitate rapid, reliable

assessment of the presence or absence of airborne powdery mildew inoculum early

in the progress of an epidemic to guide the initiation of control measures. This could

result in more reliable and cost-effective control of this disease early in the progress

of an epidemic.

Modification of the Fastprep kit protocol was necessary in order to ensure successful

and consistent amplification of target DNA. Amplification of DNA by both universal

and species-specific primers required the addition of PVP to the extraction buffer

prior to homogenization, and the addition of a phenol chloroform extraction step.

Potential PCR inhibitors (including incidental plant phenolics) were, in most cases,

assumed to be sufficiently inactivated or removed by these modifications (Porebski et

al., 1997; Boer, et al., 1995; Zhou et al., 2000). This protocol was additionally altered

for two species of powdery mildew because the DNA extraction protocol utilized by

the majority of the samples was unable to obtain amplification products using the

universal primers ITS1 and ITS4. A combination of the Fastprep plant and fungi

buffers used in conjunction with the PSS reagent followed by the described extraction

procedure resolved this problem.

Because amplification of DNA from conidia was possible by direct placement of

conidia into the PCR master mix, determining the precise level of sensitivity when

omitting the DNA extraction step when processing air samples could lead to the

16

elimination of a time consuming extraction process, potentially saving resources and

labor. Work by Williams et al. (2001), reported the sensitivity to be one- to two-fold

less when PCR was conducted directly on spores of Penicillium roqueforti. Zhou et

al (2000) reported detecting as few as two fungal spores while evaluating six different

fungi using different spore disruption methods in lieu of a DNA extraction step with

the 18s rRNA gene, a high copy number region of the genome.

Our results represent a first step toward the development of a quantitative field

detection method for E. necator. Towards this end, we have identified a promising

quantitative PCR (qPCR) primer probe combination, which is located in the vicinity of

the ITS region also employed for our standard PCR primers. Whereas, neither PCR

nor qPCR can readily distinguish between viable and dead fungal propagules, qPCR

has the advantage of being amenable to adjustments of the threshold value to

accommodate a certain amount of background from assumed dead spores that might

be present in the dormant season or in the vineyard after periods of extremely hot

weather. Through adjustment of the detection threshold for field applications, it

would likely be possible to account for residual, nonviable spores and mycelia from

prior epidemics, while maintaining the sensitivity required for the quantification of

disease pressure. Butt and Royle have described the use of spore populations as a

measure and predictor of disease severity (Krantz, 1974). Other qPCR advantages

include the added specificity of a labeled probe in addition to the two primers, as well

the higher throughput qPCR offers when several primer probe combinations

(detecting and distinguishing between different pathogens) are pooled into a single

17

reaction with different reporters in multiplex qPCR. Further research, including qPCR

and the development of molecular probes could lead to faster, more consistent and

less expensive practical field applications and in depth epidemiological studies.

The findings of this study describe a rapid, reliable, and inexpensive detection tool

that could be used in conjunction with other technologies to improve the precision of

existing risk assessment models (Gubler et al., 1996). PCR is a promising tool for

the timely detection and diagnosis of E. necator. This information may, in the future

improve the precision of existing forecasting models to better predict necessary

fungicide applications.

ACKNOWLEDGMENTS

We acknowledge the financial support of this project by the Washington Wine

Advisory Board and the Washington State University Agricultural Research Center,

We appreciate the powdery mildew samples provided by G. Saenz, F. Delmotte,

W. Mahaffee, D. Gadoury, D. Gubler, M. Miller, G. Newcomb, C. Nischwitz,

E. Bentley, and the technical support received from P. Scholberg, D. O’Gorman,

K. Bedford, and K. Eastwell. We also appreciate the support and editorial skills of

Terri Hughes and Duane Moser.

18

Fungus name fide Braun (1987) (Fungus name fide Braun & Takamatsu (2000)) Host Genus species Location Detection Blumeria. graminis (DC.) Speer Poa sp. 1 - Erysiphe. aquilegiae (Grev.) Zheng & Chen

var. ranunculi Aquilegia canadensis L. 5 - E. artemisiae Grev Tanacetum vulgare L. 1 -

E. betae (Vanha) Weltzien Beta vulgaris L. subsp. cicla

(L.) W. Koch 8 - E. cichoracearum DC var. cichoracearum. Aster sp. 1 - Coreopsis sp. 1 - Cosmos sp. 4 - Chrysanthemum maximum Cav. 1 - Taraxacum officinale Wigg. 5 - Lactuca serriola L. 1 - Rudbeckia laciniata L. 1 - E. convolvuli DC. Convolvulus arvensis L. 1 - E. cynoglossi (Wallr.) U. Braun Amsinckia tessellata Gray 3 - Pulmonaria sp. 8 - E. galeopsidis DC. Ajuga sp. 1 - E. glycines Tai Lupinus perennis L. 5 - E. liriodendroni Schw. Liriodendron tulipifera L. 11 - E. magnicellulata U. Braun var. magnicellulata Phlox sp. 2 - E. pisi DC. Medicago sativa L. 1 - Pisum sp. 3 - E. polygoni DC. Polygonum convolvulus L. 1 - Trifolium sp. 9 - E. rhododendri Kapoor Rhododendron sp. 6 - Leveillula. taurica (Lév.) Arnaud Allium cepa L. 1 - Microsphaera. alphitoides Griffon & Maubl.

(E. alphitoides (Griffon & Maubl.) U. Braun & S. Takamatsu) Quercus robur L. 6 -

M. berberidicola F.L. Tai (E. berberidicola (F.L. Tai) U. Braun & S. Takamatsu)

Mahonia aquifolium (Pursh.) Nutt. 5 -

M. euonymi-japonici Vienn.-Bourg (E. euonymi-japonici (Vienn.-Bourg) U. Braun &S. Takamatsu) Euonymus fortunei Hand.-Mazz. 2 -

M. platani (Howe) (E. Platani (Howe) U. Braun & S. Takamatsu Platanus occidentalis L. 9 -

M. Syringae (Schwein.) Magnus (E. Syringae Schwein) Syringa vulgaris L. 1 -

Ligustrum japonicum Thunb. 6 - Caragana arborescens Lam. 9 - M.nemopanthis Peck

(E. nemopanthis (Peck) U. Braun & S. Takamatsu) Ilex verticillata (L.) A. Gray 2 -

Oidium sp.a Laburnum anagyroides Medik. 7 - Podosphaera. clandestina (Wallr.: Fr.) Lév. Prunus avium (L.) L. 1 - P. leucotrica (Ell. & Ev.) Salmon Malus sylvestris Mill. 1 - Phyllactinia. guttata (Wallr:.Fr.) Lév Corylus cornuta Marsh. 6 - Sphaerotheca. aphanis (Wallr.) U. Braun

(P. aphanis (Wallr.) U. Braun & S. Takamatsu) Rubus ursinus Cham. &

Schlechtend. 7 - S. delphinii (P. Karst) S. Blumer

(P. delphinii (P. Karst) U. Braun & S. Takamatsu) Ranunculus abortivus L. 1 -

S. fusca (Fr.) S. Blumer (P. fusca (Fr.) U. Braun & N. Shishkoff) Monarda didyma L. 1 -

Cucurbita maxima Duchesne 2,8 -

19

S. macularis (Wallr.:Fr.) Lind (E. macularis (Wallr.:Fr.) U. Braun & S. Takamatsu) Humulus lupulus L. 1 -

S. pannosa (Wallr:.Fr.) Lév (P. pannosa (Wallr:.Fr.) de Bary) Rosa sp. 1 -

S. violae U. Braun (P. violae (U. Braun) U. Braun & S. Takamatsu) Viola renifolia A. Gray 8 -

Sawadaea bicornis (Wall.: Fr.) Homma Acer platanoides L. 5 - Uncinula adunca (Wallr.: Fr.) Lév

(E. adunca (Wallr. ) Fr.) Pupulus sp. 10 - U. necator (Schwein. ) Burrill

(Erysiphe necator Schwein.) Vitis vinifera L. 1,12,13,14 + Uncinuliella flexuosa Peck

(E. flexuosa (Peck) U. Braun & S. Takamatsu) Aesculus sp. 9 -

1=Prosser, WA; 2=Benton City, WA; 3=Richland, WA; 4=Kennewick, WA; 5=Pullman, WA; 6=Seattle, WA; 7=Pack Forest, WA; 8=Bellingham/Mt Vernon, WA; 9=Moscow, ID; 10=Fairbanks, AK; 11=Bent Creek, NC; 12=NY; 13=CA; 14=France

a Possibly Microsphaera guarinonii (E. communis), however, no literature describes the anamorph. Only two species were reportedby Braun on Laburnum anagyroides (M. guarinonii and L taurica.) This specimen is not a Leveillula. The USDA ARS host index (Farr, n.d.) reports a L. taurica, M. guarinonii, E. communis, and an Oidium sp. on Laburnum sp.; none of which are reported in the Americas. Braun lists E. communis as a synonym of M. guarinonii, thus three of the four species reported in the host index could be the same species according to Braun.

Table 1. Powdery mildews collected, identified and evaluated using PCR with

primers ITS1 and ITS4, and Uncin144 and Uncin511.

20

E. necator isolate Origin Source 2B17 France F. Delmotte BR8 France F. Delmotte CC43 France F. Delmotte CC12 France F. Delmotte Lat13 France F. Delmotte Be3 France F. Delmotte Turloc 1 California D. Gubler/T. Miller Orcutt 1b California D. Gubler/T. Miller Sonoma Co 1b California D. Gubler/T. Miller Fresno Co. 1e California D. Gubler/T. Miller Fresno Co. 1d California D. Gubler/T. Miller EWA E. Washington G. Grove/J. Falacy WWA W. Washington L. du Toit Mad 28 New York D. Gadoury Pal II New York D. Gadoury Fr 25 New York D. Gadoury Fr 40 New York D. Gadoury FR 38 New York D. Gadoury Mad 17 New York D. Gadoury

Table 2. List of E. necator isolates from diverse geographic origins yielding

amplification products when PCR was performed using primers specific for

E. necator.

21

FIGURES

Fig. 1. Diagram of Internal Transcribed Spacer (ITS) Region, a portion of DNA

located between the large and small ribosomal subunits. Uncin144 and Uncin511

are located between the universal primers ITS1 and ITS4.

Small subunit 18s gene Large subunit 28s gene

ITS 1 ITS 4

22

Fig. 2. Agarose gels showing amplification products from polymerase chain

reaction of internal transcribed spacer regions of selected powdery mildews using

Universal primers ITS1 and ITS4 (A and B) and E. necator-specific primer pair

Uncin144 and Uncin511 (C and D). Lane (1) Ajuga, (2) Thistle, (3) Privet, (4)

Mahonia, (5) Onion, (6) Swiss Chard, (7) Grape, (8) Violet, (9) Rose, (10) Sweet

Pea, (11) Lupine. Lanes (1) Rubus, (2) Pulmonaria, (3) Ligustrum, (4) Alfalfa, (5)

Euonymus, (6) Bioguardian spore trap stock DNA solution, (7) Bioguardian spore

trap 1:6 dilution.

23

Fig 3. Agarose gel showing amplification products from polymerase chain

reaction with E. necator conidia added directly to the master mix and using E.

necator-specific primers Uncin144 and Uncin511. Lanes (1-9) 2 conidia per PCR

reaction, (10) Positive control (extracted DNA), (11) water control.

24

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