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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
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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
___________________________________
___________________________________
___________________________________
___________________________________
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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
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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.
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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
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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.
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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
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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
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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.
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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
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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
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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
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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.
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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).
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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
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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
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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.
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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
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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.
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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.
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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.
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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
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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.
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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
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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
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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
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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
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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.
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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 -
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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.
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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.
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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
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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.
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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.
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