.. afijr.

 

“a”?! u .

1%“. a}

flu “”2“" hi“! I-

c:
1

:l-«Qu- 4':

VI

L

 

n‘

 

WIS

2 LIBRARY
L a 0% Michigan State
" University

 

 

 

This is to certify that the
thesis entitled

DISTRIBUTION, IDENTIFICATION, AND POPULATION
DIVERSITY OF ARM/LLARIA SPP. IN MICHIGAN CHERRY
ORCHARDS

presented by

MARGARET LEE ELLIS

has been accepted towards fulfillment
of the requirements for the

Master of degree in the Department of Plant
Science Fathom

 

 

7 Major Professor’s Signature

(2. N omw 100 7—

 

Date

MSU is an affinnative-action, equal-opportunity employer

 

PLACE IN RETURN BOX to remove this checkout from your record.
TO AVOID FINES return on or before date due.
MAY BE RECALLED with earlier due date if requested.

 

DATE DUE

DATE DUE

DATE DUE

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

6/07 p:IClRC/DateDue.indd-p.1

 

DISTRIBUTION, IDENTIFICATION, AND POPULATION DIVERSITY OF
ARMILLARIA SPP. IN MICHIGAN CHERRY ORCHARDS

BY

MARGARET LEE ELLIS

A THESIS
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

MASTER OF SCIENCE
Department of Plant Pathology

2007

ABSTRACT
DISTRIBUTION, IDENTIFICATION, AND POPULATION DYNAMICS OF
ARMILLARIA SPP. IN MICHIGAN CHERRY ORCHARDS
BY
MARGARET LEE ELLIS
Species of Armillaria (Fr.:Fr.) Staude cause shoestring root rot, an important

disease of Montmorency tart cherry trees in Michigan. The first objective of this study
was to evaluate the current distribution of Armillaria spp. in the cherry producing
regions. Isolates were collected from 14 counties in Michigan, and identified to species
using PCR amplification of the IGS-l region and RF LP analysis. Sexual compatibility
tests were used to distinguish between species with the same DNA banding patterns.
Isolates were identified as A. ostoyae, A. gemina, A. mellea, or A. gallica. Armillaria
ostoyae was the predominant species found on cherry, corresponding to the previous
survey by Proffer et a1. (1987). Armillaria gemina and A. gallica were found for the first
time on orchard grown Prunus spp. The second objective was to examine the population
and strain diversity of Armillaria within two orchards using somatic incompatibility tests
and microsatellites. In 2006, two tart cherry orchards in the northwest region of
Michigan were intensively surveyed. Six individual clones of A. astoyae were detected
from 22 isolates from the first orchard. From the second orchard four different species
were detected from 81 isolates. The isolates included 11 clones of A. ostoyae, four clones
of A. gemina, four clones of A. mellea, and three clones of A. gallica. These findings
expand available information on population diversity of Armillaria, providing growers

with valuable information on whether a site is suitable for orchards or vineyards.

DEDICATION

I would like to dedicate this thesis to my loving family and closest friends,
especially my father Mark Lee Ellis, mother Deborah Lynn Ellis, and my sister Elizabeth

Stewart Ellis. They have been a great source of support and guidance throughout my life.

iii

ACKNOWLEDGEMENTS

I would like to thank my major professor Dr. Ray Hammerschmidt for guiding me
in my educational and professional development in the field of Plant Pathology. I have
grown tremendously under his direction. My project in his lab will help to diversify my
knowledge and experience in this field as I continue my career. I would like to thank my
committee members, especially Dr. Tyre Proffer for his expertise in mycology, his
constant guidance, knowledge, and support. Also on my committee, I would like to thank
Dr. George Sundin for his excellent support and advice he brought to my project, and Dr.
Annemiek Schilder. This project could not have been completed without the support
from the growers, Northwest research station, and all of the extension agents. I
especially want to thank Jim Nugent for his expertise in cherry and also for the great help
in completing my survey. 1 would like to thank Dr. Shuxian Li, Dr. Glen Hartman, and
Dr. Don White from the Univeristy of Illinois for there support and inspiration to
continue a career in the field of Plant Pathology and to encourage me to continue my
education here at Michigan State University. Finally, I would like to thank the other
faculty members at Michigan State University that helped expand my knowledge in this
field, and Janette Jacobs, Samantha Hollosy, and Deanna Lanier from the
Hammerschmidt lab that extended both friendship and a helping hand in my research.

This research was funded by a grant from USDA-CSREES, who I would like to

thank for supporting this research.

iv

TABLE OF CONTENTS

 

 

 

 

 

 

 

 

 

 

 

 

 

 

PAGES
LIST OF TABLES vii
LIST OF FIGURES ....................... -- ix
KEY TO ABBREVIATIONS .......................... - -Xi
CHAPTER 1. GENERAL INTRODUTION
AND LITERATURE REVIEW__ _ _ _ 1
GENERAL INTRODUCTION 2
LITERATURE REVIEW .............. 7
Taxonomy ...... 7
Morphological features ______ 11
Life cycle 15
Control 19
Molecular methods used for species identification ,,,,,,,, 25
Somatic incompatibility 29
Distribution studies 32
The host range 36
LIST OF REFERENCES 42

 

CHAPTER 2. DISTRIBUTION, IDENTIFICATION, AND POPULATION
DIVERSITY OF ARMILLARIA SPP. IN MICHIGAN

 

 

 

 

 

 

 

 

 

 

CHERRY ORCHARDS 52
INTRODUCTION _____ 53
MATERIAL AND METHODS __ _ 57
Survey and field sampling 57
Species identification 61
DNA sequencing of the 163-] region 67
Somatic incompatibility tests ______ 67
Microsatellite tests .......................... 69
RESULTS 7]
Species identification and distribution 71
Somatic incompatibility and microsatellite results _________ 84
DISCUSSION 9]

 

LIST OF REFERENCES _

FINAL CONCLUSIONS

APPENDICES

 

APPENDIX A. Species identification results

 

APPENDIX B. Somatic incompatibility tests
and microsatellite results

APPENDIX C. Greenhouse rootstock inoculation and field trial

APPENDIX D. Glossary ......

vi

97

-_ 101

103

103

.117

122

133

LIST OF TABLES

 

 

 

 

 

 

 

TABLE PAGES
CHAPTER I

1-1 Species of Armillaria in North America with assigned NABS:

Modified from Volk and Burdsall _____ 9
1-2 Reported host range of both saprophytic and pathogenic species

of Armillaria in Michigan: Modified by Kromroy (2004) 37

CHAPTER II

2-1 Location, date, and description of sites surveyed for Armillaria

root rot 58
2-2 Tester strains used as controls to compare and identify Michigan

isolates of Armillaria to species 65
2-3 Microsatellite markers used to genetically confirm SIG’s from the

two intensively sampled orchards 70
2-4 Number of isolates collected from each county, the host, and

species distribution of Armillaria spp. in the cherry producing

regions of Michigan 76
2-5 Somatic incompatibility groups and microsatellite results for the

first orchard (site # 14), second orchard (site # 29), and seven

miscellaneous strains of A. ostoyae collected from other sites ,,,,,,,,,,,,,,,, 86

APPENDIX A

A-l Location, tree type, banding size for the IGS-l region, restriction

enzyme digest results, and species determination, for established

Armillaria isolates 103
A-2 Sexual compatibility results for strains of Armillaria identified as

A. calvescens/ A. gallica using PCR amplification of the IGS-l
region and RFLP patterns 113

vii

A-3

B-l

C-1

C-2

Sexual compatibility results for strains of Armillaria identified as
A. ostoyae/ A. gemina using PCR amplification of the 168-]
region and RFLP patterns

 

APPENDIX B

Results from somatic incompatibility test and microsatellites for
strains of A. ostoyae collected from the first orchard (site # 14)
and second orchard (site # 29) and microsatellite results for seven
A. ostoyae strains from miscellaneous sites

 

APPENDIX C

Strains of Armillaria used to inoculate grape rootstock

 

Pot setup for greenhouse inoculations for each rootstock variety ,,,,,,,,,,

viii

115

117

125

126

LIST OF FIGURES

FIGURE PAGES
CHAPTER 11

2-1 AluI banding patterns observed for Michigan Isolates of
Armillaria spp. 78

2-2 NdeI banding patterns observed for Michigan
isolates of Armillaria spp. 79

2-3 BsmI banding patterns observed for Michigan
isolates of Armillaria spp._ 80

2-4 Sexual compatibility test results for species identification of
Armillaria isolates, using haploid tester isolates 81

2-5 Sexual compatibility test results for species identification of
Armillaria isolates, using haploid tester isolates and unknown
diploid isolates 82

2-6 Distribution map of species of Armillaria in Michigan’s
cherry producing regions." 83

2-7 Somatic incompatibility tests used to define genets of
Armillaria in the first orchard (site #14) and second orchard
(site #29) 88

2-8 Distribution of A. ostoyae genets in the first orchard (site # 14)
based on results from somatic incompatibility tests and
microsatellite markers ______ 89
2-9 Distribution results of Armillaria genets for the second orchard
(site # 29) based on results from somatic incompatibility tests and
microsatellite markers 90

APPENDIX C

C-l Prepared Armillaria inoculum for the greenhouse inoculation trial
to test grape rootstocks to susceptibility to A. astoyae and A.mellea_,,_ 127

ix

C-2 Greenhouse inoculation trial to test four different grape rootstocks
,to their susceptibility to A. ostoyae and A. mellea 128

C-3 Grape rootstock field trial to test four rootstocks to susceptibility
of A. ostoyae 129

C-4 Field trial setup to test grape rootstock to susceptibility to A. ostoyae" 130

Images in this thesis are presented in color.

KEY TO ABBREVIATIONS
BS: Biological species
LSU region: nuclear large ribosomal subunit
IGS region: Intergenic spacer region
ITS region: Internal transcribed spacer region
mtDNA: Mitochondrial DNA
NABS: North American biological species
PCR: Polymerase chain reaction
rDNA: Ribosomal DNA
RAPD: random amplified polymorphic DNA
RF LP: Restriction fragment length polymorphisms
SDS-PAGE: Sodium dodecyl sulfate polyacrylamide gel electrophoresis
SI: Somatic incompatibility

SSR’s: Simple sequence repeats also known as microsatellites

xi

CHAPTER I

GENERAL INTRODUCTION AND LITERATURE REVIEW

GENERAL INTRODUCTION

The genus Armillaria (Fr.:Fr.) Staude is classified within the fungal class
homobasidiomycetes. Armillaria spp. produce gilled pileate basidiomes, typically with
an annulus around the central stipe, and grow in caespitose clumps around the base of
dead trees or stumps (reviewed in Watling et a1. 1991). The common names for this
edible fungus, honey mushroom or honey agan'c, come from the pileus or cap of the
mushrooms that are tan/amber/honey colored. The other common name given to
Armillaria is the oak fungus because of the regular occurrence of one of the major
species (Armillaria mellea (Vahl:Fr.) Kummer) on this forest host. Armillaria spp. are
among the few fungi known to produce rhizomorphs. These shoestring-sized, black or
brown root-like strands can frequently be found under the bark of dead forest trees, along
decorticated logs on the forest floor, and in the soil around infested trees (IPM:
Landscape and Turf: Armillaria Root Rot of Trees and Shrubs 2000).

In forest ecosystems, Armillaria generally functions as a saprophyte and
decomposer of fallen logs. In this role, Armillaria is a white rot fungus, breaking down
both the lignin and cellulose components of wood (reviewed in Kile et a1. 1991).
However, Armillaria can also act as an opportunistic pathogen on'stressed or declining
trees, or as a primary and virulent pathogen (Shaw and Roth 1978). As a pathogen,
Armillaria causes a soilbome disease, using rhizomorph growth from an established food
source to spread the fungus through the soil from tree to tree (Jones and Sutton 1996).
Once the rhizomorphs penetrate through the root surface, the mycelium of the fungus
spreads out across the vascular cambium and phloem forming white mycelial fans, a

diagnostic feature of Armillaria infection. The destruction of phloem in the cambium

causes tree death (reviewed in Morrison, Williams, and Whitney 1991). The role of
Armillaria and its pathogenicity depends on the host, environment, and species of
Armillaria present (reviewed in Gregory et al. 1991).

As a pathogen, species of Armillaria infect a broad range of trees, shrubs, vines,
and some herbaceous plants. The disease caused by Armillaria is often referred to as
Armillaria root rot or shoestring root rot because it produces rhizomorphs (IPM:
Landscape and Turf: Armillaria Root Rot of Trees and Shrubs 2000). In Michigan,
Armillaria spp. are the causal agents of a serious localized disease in tart (sour) cherry
(Prunus cerasus) orchards. These pathogens infect other fruit orchards, forest, tree
stands, landscapes, and Christmas tree plantations (Banik et a1. 1995; Proffer et al. 1987;
Smith et. al. 1990, 1992, 1994).

In 1987, Proffer et al. surveyed the distribution of species of Armillaria in tart
cherry orchards in Michigan and found that Armillaria ostoyae (Romagn.) Herink was
the predominant species in the Northwestern orchards, with A. mellea becoming more
prevalent further south in the state. Most of the orchards surveyed by Proffer et al.
(1987) are now out of cherry production because there is no consistently effective control
for Armillaria root rot disease. According to Louvet (1979) fungicide injections are not
successful because the movement of systemic fungicides is typically acropetal. In order
for fimgicides to be affective in controlling Armillaria, translocation must be basipetal.
Kissler et al. (1973) found soil fumigation effective in destroying root fiagments infected
with Armillaria in orchards, vineyards, and floriculture operations (reviewed in Hagle
and Shaw 1991 ). Although fumigation practices have been able to show improvement in

infected sites, they do not provide complete control due to the depth that Armillaria can

be found in the soil (Bliss 1951) and also because the bark acts as a protective layer
preventing penetration of chemicals into infected tissue that remains in the soil (reviewed
in Hagle and Shaw 1991). Another restriction to using fumigants is that they can not be
applied in established orchards or forest stands. Not growing susceptible crops, such as
grasses, can reduce inoculum; however, infected roots can remain in the soil for several
years, even decades according to James Nugent, 2005, personal communication, (District
Horticulturist and coordinator of the Northwest Michigan Horticultural Research Station,
Traverse City, MI 49684) and Munnecke et al. 1981. Unless all of the infected roots are
removed from the field there is still a high potential for later infection. T richoderma
viride Pers.:Fr., Scytalidium lignicola Pesante, Peniophora gigantea (F r.) Massee,
Rhizoctonia Iamellifera Small, Pleurotus ostreatus (Jacq.:Fr.) P. Kumm., Coriolus
veriscolor (L.:F r.) Quél, Stereum hisutum (Willd.:F r.) S.F. Gray, Xylaria hypoxylon
(L.:F r.) Grev., and Hypholomafascicular (Huds.:Fr.) Kumm. are just some of the fungal
species shown to be antagonistic against Armillaria and have been tested as potential
biological controls. One nematode, Aphelenchus avenae, was found to reduce fungal
growth and vigor in a French vineyard. The genus T richoderma Pers.:F r. has been the
most studied group of fungi for biological control since they are common soil inhabitants
(reviewed in Hagle and Shaw 1991). However, according to Munnecke et al. (1981)
biological controls are hard to establish because the host and the biological control agent
interaction must happen in a relatively short time under rather specific environmental
conditions in order to be successful.

Because Armillaria continues to be a problem for Michigan cherry growers and

new sites have been reported to be infected that were previously not surveyed by Proffer

et al. (1987) a reevaluation of current orchards for the distribution of species of
Armillaria is needed. The technique Proffer et al. (1987) used relied on sexual
compatibility tests first used by Hintikka (1973), using single basidiospore isolates
collected from basidiomes. Since the 1987 survey, new DNA-based techniques allowing
identification using fimgal tissue from rhizomorphs from roots and soil, or mycelia from
fans found under the bark of diseased trees have been published (Harrington and
Wingfield 1995). Since the methodology used by Proffer et al. (1987) relied on
basidiomes, species not producing basidiomes during their survey period would have
been missed.

The next step after determining the current species of Armillaria in orchards
would be to examine the clonal spread with in an orchard site. The information gathered
from this study would help in the understanding of populations of Armillaria in Michigan
affecting the cherry orchards today. Somatic incompatibility tests and microsatellite
markers or simple sequence repeats (SSRs) are currently used to identify individual
clones of Armillaria. Somatic incompatibility tests involve pairing test isolates on a
culture medium and examining the visible interaction of the expanding colonies as they
make contact. The formation of a barrage zone along the line of contact indicates the
isolates are different clones. Microsatellite markers are used to confirm genetic variation
between individual isolates from different clones.

Many cherry orchard sites in Michigan are being converted into grape vineyards
due to the higher value of the crop per acre. New growers, not knowing the history of the
site and some established growers converting their orchard lands are often unaware that

Armillaria has been reported on grapes (Vitis spp.) in Europe, Australia, Brazil, and

California (Hood et al. 1991). Michigan’s grape industry is much younger than those of
both Europe and California, and the effects or presence of infection with Armillaria have
not yet been reported. Michigan does have one of the species of Armillaria reported on
grapes, A. mellea and another pathogenic species A. ostoyae that could be a potential
problem. In the Northwestern cherry-producing region where many of the vineyards are
currently being established, A. ostoyae tends to be the predominant species and it is
known as a virulent pathogen with a wide host range. While most reports of A. ostoyae
are on conifers, it has proven to be an aggressive pathogen of cherry in Northwest
Michigan, as seen by Proffer et al. in the 1987 survey. With the increase of viticulture in
the state of Michigan it is important to assess A. astoyae and A. mellea for their ability to
infect grape rootstocks used in Michigan.

The hypotheses of this research were: (1) that new species of Armillaria would be
identified on cherry using molecular identification techniques not reliant on basidiomes
as in the 1987 survey by Proffer et al. (1987); (2) the diversity of species in an orchard
may be much greater than previously reported; and (3) that the current Armillaria species
known to infecting orchard grown Prunus will move to grape when orchards are
converted to vineyards. To test these hypotheses a broad distribution survey of orchards
in the cherry producing region of Michigan was done, as well as a clonal mapping of
Armillaria within two Montrnorency tart cherry orchards using somatic incompatibility
tests and microsatellite markers. The third hypothesis was tested by beginning
greenhouse and field trial inoculations of grape rootstocks grown in Michigan for the

future assessment of susceptibility to A. ostoyae and A. mellea.

LITERATURE REVIEW:
TAXONOMY

The taxonomic and nomenclatural status of the genus Armillaria (F r.:Fr.) Staude
has gone through many changes over the last century. The first reports of mushrooms
that would now be classified as Armillaria are believed to be from either 1729 by Micheli
or 1755 by Battarra. Fries (1819) first established the genus Armillaria as a tribe of
Agaricus, which included Agaricus melleus Vahl:Fr., now classified as Armillaria mellea,
and later treated it in the Systema Mycologicum (Fries 1821). In 1825 Fries (1825) placed
A garicus melleus in the tribe Lepiota, abandoning the tribe Armillaria but later reversed
himself in 1938 (Fries 1838) by replacing it into the tribe Armillaria. In 1957 Staude
raised Armillaria to generic rank, including four species; Agaricus mucidus, Ag. melleus,
Ag. aurantius, and Ag. robustus. Today Ag. aurantius and Ag. robustus are considered to
be in the genus T richoloma (Fr.) Staude, and Ag. mucidus is placed in the genus
Oudemansiella Spegazzini (or Mucidula Pat.). However, Singer (1951, 1955, 1986)
believed that Staude’s entry was inadmissible according to the International Code of
Botanical Nomenclature. Singer argued that it was Kummer (1871) who should be given
credit as the author. In reviewing the nomenclatural trail, however, both Donk (1949,
1962) and Watling et al. (1982) supported Staude as the valid author for the genus
Armillaria (reviewed in Volk and Burdsall; Watling et al. 1991).

The nomenclatural confusion was compounded by Karsten (1881) Who created
the genus Armillariella in 1881. Some authors, following the lead of Singer, continued to
use this genus name into the 1970’s. Karsten, however used Ag. melleus as the type
species which was within Fries’ tribe Armillaria. The use of the genus name

Armillariella was ended afier the taxonomic review by Watling, Kile, and Gregory in

1982, which established Armillariella as an obligate synonym of Armillaria (reviewed in
Volk and Burdsall; Watling et al. 1991).

Prior to the mid-seventies, Armillaria root rot was considered to be caused by a
single polymorphic species, Armillaria mellea sensu Iato. Studies examining the
bifactorial sexual incompatibility system in Armillaria changed this view. Hintikka
(1973) developed a technique of pairing single basidiospore isolates on Petri plates to
demonstrate sexual compatibility. Additional work done by Korhonen (1978) helped to
distinguish five biological species (BS) of Armillaria in Europe (BS A = A. borealis
Marxmiiller & Korhonen, BS B = A. cepistipes Velenovsky, BS C = A. ostoyae, BS D =
A. mellea, BS E = A. gallica Marxmiiller & Romagnesi (a.k.a. A. bulbosa (Barla) Kile &
Watling and A. lutea Gillet). Today, Armillaria is divided into around 40 distinct
biological species and is found on every continent, with the exception of Antarctica,
infecting a huge range of hosts (reviewed in Volk and Burdsall).

In North America, the genus Armillaria was‘divided into 10 North American
Biological Species (NABS) by Anderson and Ullrich (1979) using the Hintikka (1973)
technique as expanded by Korhonen (1978). Most of the North American biological
species (NABS) of Armillaria, identified using the sexual compatibility system, have
been formally equated to European species or described as new species. Assigned NABS
Roman numerals designate Armillaria species with the exception of A. tabescens (Scop.)

Emel which has not yet been assigned a number (Table 1-1).

Table 1-1: Species of Armillaria in North America with assigned NABS:

Modified from Volk and Burdsall

 

Ecological
Role/
Pathogencity

Author of

NABS Species Author Synonyms Synonyms Distribution

 

Pacific
Northwest and
Northern United
States found
most often on

Armillaria
ostoyae

(Romagn.)
Herink

Armillaria
obscura

(Schaeff)

Herink [nomen

ambiwm]

conifers in the
northern conifer
zone

Pathogenic

 

II

Armillaria
gemina

Bérubé &
Dessureault

Limited to the
Eastern United
States and
Southeastern
Canada on
hardwoods

Generally not

considered
pathogenic

 

III

A rmillaria
calvescens

Bérubé &
Dessureault

Northeast of the
United States
and the
Southeast in
Canada on
maples and
hardwoods

Saprophyte in
the soil but can

actas an

opportunistic

pathoggn

 

Armillaria
sinapina

Bérubé &
Dessureault

Northeast United
States mostly on
hardwoods and
in the Pacific
Northwest on
conifers

Pathogenic

 

VI

A rmillaria
mellea

(Vahl:Fr.)
Kummer

Eastern United
States and
Canada, and
Northern
California on
hardwoods and
occasionally
conifers in
mixed forests

PathogeLiC

 

 

VII

 

Armillaria
gallica

 

Marxmuller
& Romagn

Armillaria
Iutea

Gillet [nomen

ambiguum]

 

Armillaria

 

bulbosa

 

(Barla) Kile &

Watling
[misapplied
name]

 

Eastern and
Southern United
States and
Midwest on
hardwoods

 

Saprophyte
found in the
forest soil

 

 

 

Continuation of Table 1-1: Species of Armillaria in North America with assigned NABS:

Modified from Volk and Burdsall

 

 

 

 

 

Ecological
Author of Role/
NABS Species Author Synoans Synonyms Distribution PathoLencity
Pacific
Armillaria Volk & Northwest on
IX nabsnona Burdsall hardwoods
Found only in
Idaho and
British
X Unnamed Columbia
probably
Armillaria Morrison et al. Pacific
XI cepistipes Velenovsky Species F 1985 Northwest
Eastern United
States, Midwest,
and in the West
in Texas and
Oklahoma.
Armillaria (Scop.) Common on
tabescens Emel Oak. Pathogenic

 

 

 

 

 

 

 

10

 

MORPHOLOGICAL FEATURES

The basidiomes of Armillaria are generally found in caespitose clumps (A.
ostoyae, A. gemina, A. mellea, and A. tabescens) at the base of a tree or stump, in
gregarious clusters (A. calvescens, A. gallica, and A. nabsnona), in clumps of two to three
or single (A. sinapina). The pileus is fleshy and the color of the pileus can be honey
colored (A. mellea), brown to brownish-yellow (A. ostoyae, A. gemina, A. sinapina, A.
calvescens, and A. gallica), tan (A. calvescens, and A. gallica), pinkish-brown (A.
calvescens, and A. gallica), ivory (A. tabescens), or orange brown (A. nabsnona). The
pileus is usually smooth (A. calvescens, A. mellea, A. sinapina, A. gallica, A. nabsnona,
and NAABS XI) but can be scaly (A. ostoyae, and A. gemina). The flesh of the pileus is
pale, while the stipe is white at first but then can turn, brown, tan, or yellow. The stipe is
central and typically annulate, with the exception of A. tabescens, with floccose-
membranous to arachnoid veil. The annulus can be thick and wooly (A. mellea), thick (A.
ostoyae, and A. gemina), or cortinaceous (arachnoid, cobwebby) (A. calvescens, A.
sinapina, A. gallica, and NABS XI). The gills are often decurrent, but can be sinuate,
adnexed, or subdecurrent depending on the species. The basidia are usually 4-spored but
can be 2-spored and thin-walled, often with a basal clamp-connection. The spores are
often thick-walled and hyaline to cream colored and produce a white to cream-colored
spore print (reviewed in Volk; Watling et al. 1991).

Armillaria was first physically described as a gilled mushroom or basidiome in
Fries Systema Mycologicum (1821). The ontogeny of Armillaria and other agaricus
fungi was first described by Hoffman (1861). Basidiome development of Armillaria spp.

was also studied by Hartig (1874), Beer (1911), and Atkinson (1914). Production of a

11

basidiome in culture was first described by Molisch (1904) where he used autoclaved
bread to grow the fungus. Armillaria mellea was grown from basidiospores to basidiome
by F alck (1907). F alck reported that light was necessary for basidiome development.
Since Molisch and Falck there have been many subsequent studies reporting development
of basidiomes in vitro (reviewed in Garraway et al. 1991). Although many reports show
success in development of basidiomes in vitro, the techniques are not reliable and there
are many limitations as described by Ullrich and Anderson (1978). The in vitro
development of basidiomes is also variable depending on the species (reviewed in
Garraway et al. 1991).

Rhizomorphs are another morphological structure that characterizes the genus
Armillaria. Rhizomorphs are defined as discrete, filamentous aggregations that are
highly differentiated, fully autonomous, and grow apically. They are more complexly
organized than mycelial cords, often produced by fungi which grow on the forest floor.
Mycelial cords are aggregations of parallel, undifferentiated hyphae. Rhizomorphs are so
distinctive that they were first described as a separate species, Rhizomorphafi'agilis Roth.
Schmitz (1848) published the description of different forms of R. flagilis, including R.
subterranean found in the forest soil and R. subcorticalis found growing. beneath the bark
of trees. Robert Harti g (1874) later provided proof that rhizomorphs belonged to the
mushrooms of the genus Armillaria (reviewed in Garraway et al. 1991).

Rhizomorph organization is described in detail by Hartig (1870, 1874). In 1877,
the apical growth of rhizomorphs was described by Brefeld, and in 1969 Motta improved
the descriptions with the use of electron microscopy. Harti g described the thallus

organization of rhizomorphs as having three layers including; a cortex, subcortex, and

12

medulla layer. The three layers can be characterized by three different types of hyphae:
skeletal, binding, and generative. The mucilaginous nature of the tip of the rhizomorph
and the differences in the cell wall layers were also described by Hartig. The cortex layer
is composed of melanized densely packed cells. Bloomfield and Alexander (1967), and
Khuo and Alexander (1967) found that the melanin content of the cortex protects the
inner layers of rhizomorphs from potential hazards in the environment. The subcortex is
a transition layer from the cortex to the medulla where the hypha becomes wider in
diameter and looser. The medulla layer is composed of loose hyphae and forms a central
canal. J enning (1984) described how the medulla layer is responsible for both water and
nutrient transport and Smith and Griffin (1971) described how the canal is responsible for
oxygen translocation (reviewed in Garraway et al. 1991).

The nutrient uptake, transport, and requirements of rhizomorphs have been
studied. Falck (1912) was the first to propose the idea that rhizomorphs were used for
transport of nutrients and water. Chlorides, phosphates, and ammonium ions were all
shown to transport through rhizomorphs by Morrison (1975), later confirmed by
Anderson and Ullrich (1982) using radioactive labeling. Eamus and Jennings (1984)
found a considerable gradient of water and turgor potential from the tip to the base of
rhizomorphs, and hypothesized that long-distance transport occurred predominantly by
solutes moving along vessel hyphae of the medulla. This view was later supported when
Eamus and Jennings (1985) measured the internal structure and hydraulic conductivity of
rhizomorphs (reviewed in Garraway et al. 1991).

There have been many in vitro studies on the nutrient requirements of Armillaria.

The molecules: carbohydrates, lipids, phenols, and alcohols have been examined.

13

Addition of low molecular weight alcohols to glucose media were shown to increase the
growth rate of A. mellea (Vance and Garraway 1973). Bell (1970), Cruickshank and
Perrin (1971), and Masuko et al. (1970) have shown phenols to be inhibitory to many
fungi (reviewed in Vance and Garraway 1973). However, when low molecular weight
alcohols such as ethanol, propanol, butanol, or isobutanol were added to media, the
inhibitory affect of phenols on the growth of Armillaria isolates decreased (Vance and
Garraway 1973). Studies have suggested many reasons why phenols inhibit growth.
Smith and Griffin (1971) suggested that in A. luteobubalina Watling & Kile (previously
reported as Armillariella elegans) polymers formed by phenols which oxidize to
melanins and lignins can be incorporated into cell walls of rhizomorphs causing increased
rigidity and thus prevent cell wall expansion. It has also been suggested that phenols
inhibit enzymes that could be important in mycelial grth and rhizomorph
morphogenesis. Phosphorylases (Schwimmer 1958), cellulases (Kosuge 1969),
transaminases (Braunstein 1947), and decarboxylases (Hartman et al. 1955) are all known
to be inhibited by phenols (reviewed Vance and Garraway 1973).

In pathogenic species of Armillaria, rhizomorphs are used as a mode of infection.
The growth habit of rhizomorphs in different species of Armillaria is correlated with
pathogenicity (Morrison 1989, 2004). Morrison examined 15 species of Armillaria from
Europe, North America, Australia, and New Zealand; species whose rhizomorphs
branched dichotomously were more virulent then those that branched monopodially
(Morrison 2004). Rhizomorphs infect their host by sending branches into their host as
they grow epiphytically along the host (Garrett 1956). Once infected, the root acts as the

inoculum or food-source for further spread. Such infected sources are important in the

14

success of progressively establishing infections. Garrett, using woody inoculum of
varying sizes with potato (Solanum tuberosum L.) tubers as the potential host at different
distances from the inoculum demonstrated that the successful establishment of new
infections depended upon factors such as the size of the food-base and the distance from

the food-base (Garrett 1956).

LIFE CYCLE

Armillaria has a unique life cycle compared to many other basidiomycetes. The
initial interactions of a compatible mating in most basidiomycetes result in cell fusion
(plasmogarny) with delayed nuclear fusion (karyogarny). A dikaryotic stage (n+n) is the
predominant nuclear condition in the vegetative cells of most basidiomycetes (Peabody et
al. 2000). Clamp connections, a characteristic found in most basidiomycetes, allow for
nuclear migration where a dikaryotic cell donates one nucleus to a new monokaryotic
cell. This process of nuclear migration and mating of many basidiomycetes was first
examined by Buller (1931) with Coprinus Pres. and is now known as the Buller
Phenomenon. This process can be used to distinguish biological species, and involves
pairing of a dikaryotic and monokaryotic isolate. If the cells from the dikaryotic isolate
are able to successfirlly transfer nuclei to the cells of the monokaryotic isolate, which is
“dikaryotized’, the isolates are said to be from the same biological species and are
compatible. Today this unique phenomenon is used for many basidiomycetes to
distinguish biological species within a genus (reviewed in Callac et al. 2006; Rizzo and

May 1994).

15

In contrast to most basidiomycetes, the predominant nuclear stage of Armillaria is
diploid. The basidiomes, mycelial fans found under the bark of trees, and rhizomorphs
tend to be diploid. Haploid mycelium is rare but has been seen in nature (Rishbeth 1985).
The dikaryotic stage of Armillaria is limited to a short period right after the haploid
mycelium of two different mating types mates fuse. Karyogamy occurs immediately and
the hyphae become diploid. However, some species of Armillaria appear to have a
second dikaryotic stage in the subhymenial cells that form clamp connections with young
basidia. This was first observed by Korhonen (1980) in Armillaria ostoyae, which helped
to explain why clamp connections were often present in fruiting bodies of some species
of Armillaria. Korhonen (1980) observed that fruiting bodies of Armillaria ostoyae
produced in the lab were clampless unlike ones seen in nature where the basidia have a
basal clamp. From this initial work, two hypotheses were developed to explain how this
could happen. The first hypothesis is that Armillaria goes through two diploidization and
two haploidization cycles prior to formation of the basidia (Grillo et a1. 2000). Several
earlier works favored this hypothesis, and fluorescence microspectrophotometric work
supports this theory (Korhonen 1980; Peabody et al. 1978; Peabody and Peabody 1984,
1985, 1986, 1987; Tommerup and Broadbent 1975). Another theory is that Armillaria is
a genetic mosaic. This theory can be supported by Peabody et a1. (2000) who first found
haploid, monokaryotic fruiting bodies occurring in nature and were later able to
demonstrate that the basidiomes of A. gallica were a genetic mosaic, by comparing
genotypes of different cells within the same fruiting body. Furthermore, Rizzo and May
(1994) were able to produce 2N+ N mycelium using A. ostoyae strains in the lab, and

suggested that this could be a stable phase in nature. If this stage was stable in nature and

16

basidiomes were produced from this mycelium it could explain diversity of allelic
combinations, thus supporting the genetic mosaic theory. Peabody and Peabody (2003)
support both hypotheses, stating that because of genetic mosaicism in the basidiome, A.
gallica has a life cycle with two diploidizations and two haploidizations. Like other
basidiomycetes, Armillaria retains the ability to mate diploid mycelium with compatible
haploid mycelium to produce new diploid cells (Anderson and Ullrich 1982; Korhonen
1978; Rizzo and Harrington 1992). Diploid-haploid mating along with haploid-haploid
mating is used today to identify biological species.

Armillaria root rot infections involve a source of inoculum, such as colonized
stumps or roots from a previously infected host. From the pre-established inoculum
source, rhizomorphs can grow through the soil or along a root, thus acting as the primary
mode of dispersal. Rhizomorphs are not the only means of infection; rhizomorph
infection does not occur in some species such as A. tabescens. Armillaria tabescens
infection occurs via root to root contact or interlocking root systems. Basidiospores can
also act as a source of inoculum by colonizing a stump. Stump colonization by
basidiospores is believed to be rare but can play an important role in establishing new
foci or infection centers of disease (Rishbeth 1970). Genotype identification studies have
shown some indirect evidence of spore infections. Many researchers have tried to
replicate this process and have had either no success or limited success infecting stumps
with basidiospores (reviewed in Redfem and Filip 1991).

Once in contact with a new host invasion of this host can happen one of three
ways: from pre-exiting lesions in which the pathogen was formerly held in check by host

resistance, from an epiphytic position on a root, or from outside by newly arrived

17

rhizomorphs (reviewed in Redfem and Filip 1991). Rhizomorph infection begins with its
attachment to a root and the hardening of the mucilaginous substance at its tip. Once this
occurs a single hypha develops from the rhizomorph’s tip, which penetrates the outer
layer of cork cells by mechanical force. After initial penetration, the rhizomorph
branches spread out laterally and radially into the bark. Toxins are also believed to be
produced by the fungus, helping host colonization. Shallow brown spots on the bark’s
outer parenchyma and cork layers are sloughed off and the fungus eventually reaches the
cambium and cankers develop. Once the fungus reaches the cambium, mycelial fans can
form penetrating the cambium. Bark tissue becomes necrotic (reviewed in Morrison et
al. 1991).

Disease and symptom development upon infection depend greatly on the host and
species of Armillaria. Symptoms of Armillaria infection include: reduction in shoot
growth, defoliation, crown dieback, stress-induced reproduction, canker development at
or above the root collar, and resin exudation. Because many of the symptoms are non-
specific to Armillaria, the presence of basidiomes, rhizomorphs, mycelial fans, and
pseudosclerotial plate (zones line) formation are important indicators to confirm
Armillaria as the cause of disease. Dead trees in a circular pattern with new. growth
coming back up in the center are another indicator of Armillaria root disease, both in the
field and in forests. The circular pattern is due to the radial expansion of the pathogen,
with the most recent death at the edges of the pathogens growth.

How the disease and corresponding symptoms develop is not well understood.
Two possible mechanisms have been proposed. One is that the disruption of the host’s

vascular system by the fungus and the host’s response to infection is the cause of the

18

observed symptoms. The explanation is supported on the observations of shoot decline
and changes in foliage seen over time due to infection by Armillaria and the destruction
of the host’s vascular system. A second proposed mechanism is that toxins produced by
Armillaria cause symptoms to develop (reviewed in Morrison et al. 1991). This has been
observed in conifers, infected with root rot pathogens Armillaria and Heterobasidion

annosum (F r.) Bref.(Rishbeth 1985).

CONTROL

Control of Armillaria root rot has been proved difficult. Many cultural and
chemical controls have been investigated and none have been highly successful or easy to
implement (Turner and Fox 1988). The control of Armillaria root rot depends on the host
and the environment in which one is trying to control the disease, so before control
measures can be taken these factors need to be assessed to try and find the most effective
control.

In forests and orchards, the use of resistant species or rootstocks, avoidance of
hazardous sites, cultural manipulations, chemical applications, and biological methods
have all been assessed for control of Amrillaria root rot. Currently the best method of
management seems to be a combination of methods: avoiding areas with high disease
hazard, removal of infected stumps, and the reduction of other inoculum sources such as
roots (reviewed Shaw and Roth 1978; and Hagle and Shaw 1991).

In California orchards and vineyards, fumigation or heating, along with biological
controls have provided temporary control (Munnecke 1973, 1976, 1981). The control is

temporary because the fumigant does not penetrate deeply enough into the soil (Bliss

19

1951). This has been observed in vineyards where following fumigation the newly
established vines are healthy for several years, but once the vine’s roots grew deep
enough they came in contact with inoculum that escaped fumigation and were
subsequently infected (Gubler 1992).

Host resistance is an option that has been found to be somewhat effective in
managing Armillaria root rot in both forests and orchards. Morrison (1981) and Hadfield
et al. (1986) found that ponderosa pine (Pinus ponderosa Dougl. ex Laws.) is much more
resistant than true firs (Abies spp.) and Douglas-fir (Pseudotsuga menziesii (Mirb.)) in
Western North America. Greig and Strouts (1983) found Douglas-fir was more resistant
than Scots pine in Great Britain. Resistance of fruit rootstocks to Armillaria species has
also been tested. Variation in citrus (Citrus spp.) rootstock susceptibility to Armillaria
root rot was observed by Thomas et a1. (1948). Guillaumin et al. (1989) observed plum
(Prunus spp.) to be more resistant than almond (Prunus amygdalus Batsch.), apricot
(Prunus armem'aca L.), cherry (Prunus spp.), and peach (Prunus persica Sieb. & Zucc.)
which were highly susceptible to Armillaria mellea in France (reviewed in Hagle and
Shaw 1991). In Michigan, cherry rootstocks were tested in cold frames; P. mahaleb L.
(mahaleb) rootstock, commonly used with tart cherry, was more susceptible than P.
avium L. (mazzard) rootstock, commonly used with sweet cherry Prunus avium L.
(Proffer et al 1988). However, this resistance may not hold up in the field, as seen by
James Nugent. According to Nugent, the death of trees grafted on mazzard rootstock
does not occur as quickly, but the final mortality appears to be the same. In other words,

trees grafied on mahaleb rootstock planted into inoculated soil often die in two to four

20

years, whereas trees on mazzard rootstock die in four to seven years (N ugent, 2005,
personal communication).

Reduction of inoculum is another control option. This involves the physical
process of turning the soil to a considerable depth and removing all the stumps and roots
of infected trees, and then cropping the land for several years with non-woody plants
(reviewed Hagle and Shaw 1991). In orchards in Michigan, this method is time
consuming and requires many years of abandonment of that orchard site. It also does not
ensure all of the removal of inoculum due to the depth of the root systems. It only takes
one piece of infected root tissue to begin new infection, and the inoculum can survive for
several years. For these reasons, this control method is often not feasible or reliable to
cherry growers with heavily infected orchards (N ugent, 2005, personal communication).

Other cultural controls, such as root collar excavation, and digging trenches to
reduce spread have also been tested. In New Zealand, trenches were dug and lined with
plastic and then back filled to serve as mechanical barriers to rhizomorph and root spread
in kiwifruit (Actinidia spp.) orchards. However, this method was not cost effective due to
the lack of profitability of kiwifruit (reviewed in Hagle and Shaw 1991). The efficacy of
root collar excavation of grape, as a post-infection control method, was examined by
Baumgartner (2004). In this study, root collars were exposed by removing the soil in an
area of approximately 0.5 m in diameter around the root collar, and 0.3 m in depth. The
goal was to determine if over time the mycelial fans would recede before severe disease
occurred resulting in decay of vascular tissue. Baumgartner (2004) found that in one
grape vineyard, excavation significantly increased yield and cluster weight of

symptomatic grapevines. Also from this study it was revealed that mycelial fans had

21

receded from root collars of symptomatic-excavated grapevines when reexamined the
following year (Baumgartner 2004).

Chemical fumigants, such as methyl bromide, carbon disulphide, and chloropicrin
have been investigated as a means of control (reviewed in Hagle and Shaw 1991).
Methyl bromide has been recommended as a control for Armillaria root rot (Filip and
Roth 1977; and Munnecke et al. 1973, 1981). The use of carbon disulfide as a soil
fumigant was first employed in Europe by Girard (1894) and Oberlin (1984) and was
later recommended by Home (1914) for destruction of A. mellea in California (reviewed
in Bliss 1951). Bliss (1951) looked at nine different soil fumigants including carbon
disulfide, formalin, tetrachlorethane, chloropicrin, chlorine, sulfur dioxide, ammonium
hydroxide, ethylene oxide, and dichlorethyl ether. Bliss (1951) extensively examined
carbon disulfide as a fumigant in citrus soils and found that injections at regular intervals
over infected sites after the removal of stumps protected fruit crops in California from
infection by Armillaria (Bliss 1951). The use of methyl bromide and carbon disulfide
has been shown to predispose the pathogen to attack by antagonistic soilbome fungi, such
as T richoderma viride Pers.:Fr., helping to eradicate mycelium in buried wood segments.
However, this method is not effective in killing all of the Armillaria in infected roots
because the fumigants do not penetrate deeply enough into the soil (reviewed in
Baumgartner 2004). Chemical treatments including ammonium sulphamate, 2,4,6-T
(2,4,5—trichlorophenoxyacetic acid), Vorlex (80% chlorinated hydrocarbons, 20% methyl
isothiocyanate), Picfume (99% chloropicrin), carbon disulfide, Dowfume (98% methyl
bromide), and Vapam (32.7% sodium N-methyldithiocarbamate) have also been tested to

reduce inoculum in forests by fumigation of infected stumps (Filip and Roth 1977; and

22

Rishbeth 1976). More recently Aguin et al. (2006) studied the in vitro capability of four
sterol demethylation inhibitors including cyproconazole, hexaconazole, propiconazole,
and tetraconazole, and another six downwardly mobile systemic chemicals including,
azoxystrobin, cubiet (copper bis(ethoxy-dihydroxy-diethylamino)sulfate), fosetyl-Al,
potassium phosphite, sodium tetrathiocarbonate, and 2-(thiocyanomethylthio)
benzothiazole for possible control of A. mellea. Their results indicated that the four sterol
demethylation inhibitor fungicides were the best at inhibiting fungal growth, especially
cyproconazole and hexaconazole which showed a reduction of 67-72% at an ECSO dose of
only 1 mg AI litre'l (Aguin et al. 2006).

Fungicides have not provided long term control in the field. Delivering the
fungicide to the infected tissue is a difficult task because the movement of systemic
fungicides is typically acropetal, and when it does move into the root system it may not
reach all of the infection (reviewed in Hagle and Shaw 1991). Adaskaveg et al. (1999)
observed that the fungicide propiconazole when injected into almond trees was effective
at increasing life of the trees for about two years before tree death, whereas most of the
control trees at similar stages of decline at the time of treatment died within four months
(Adaskaveg et al. 1999). An earlier study by Turner and Fox (1988) looked at the
potential of ergosterol-biosythesis inhibitors (EBI’s) in vitro, including: hexaconazole,
flutriafol, and fenpropidin and also at a guanide, a phenolic, and a mixture of cresylic
acids for control to A. mellea, A. ostoyae, and A. gallica (published under the synonym A.
bulbosa). At a concentration of 500 mg ai/ml, all of the fungicides were effective in
controlling Armillaria root rot. The EBI’s as well as the gaunide showed effectiveness at

lower concentrations in controlling of Armillaria root rot (Turner and Fox 1988).

23

Another fungicide, Armillatox, a phenolic emulsion, has been specifically marketed for
control of Armillaria root rot. This fungicide was developed after successful control of
Armillaria with the compound creosote (Bray 1970). However, Pawsey (1973) found
creosote to be phytotoxic, and doubted its ability to control Armillaria (reviewed in Hagle
and Shaw 1991). Redfem (1971) applied Annillatox to both forest and agricultural soils
and found that the fungicide reduced rhizomorph growth but only at the highest
concentration in agricultural soils. In results he concluded that there were no beneficial
effects from using the fungicide (Redfern 1971).

Biological controls have also been found to be partially successful, especially
when implemented with other control methods such as fumigation or heat as observed by
Bliss (1951) and Munnecke (1976, 1981). The most studied biological controls for
Armillaria are species of T richoderma. Two fungitoxic compounds from T richoderma
species have been isolated including trichoderrnin and an unidentified compound by
Ishikawa et al. (1976). Both in vitro tests and field trials have demonstrated the
antagonistic effects of T richoderma species, especially T. viride. Other fungi have also
been looked at for possible antagonistic effects to Armillaria such as Penicillium
Link:Fr., Peniophora Cooke, Scytalidium Pesante, Rhizoctonia Iamellifera, Pleurotus
ostreatus, Coriolus veriscolor, Stereum hisutum, Xylaria hypoxylon, and Hypholoma Fr.
Quél. Sokolov (1964) showed that Penicillium and Peniophora were antagonistic in in
vitro tests when plated with Armillaria. Cusson and LaChance (1974) were able to show
that Scytalidium Iigm'cola Pesante produced a toxin, syctalidin, with antifungal properties
toward Armillaria. F edorov and Bobko (1989) found that Peniophora gigantae and

Pleurotus ostreatus were effective in preventing Armillaria from colonizing fresh cut

24

stumps. Redfem (1969) found Hypholoma, a cord-forming fungus, to be competitive
with Armillaria in colonizing stumps. One nematode, Aphelenchus avenae was found to
reduce fimgal growth and vigor in a French vineyard. (reviewed in Hagle and Shaw
1991). However, although many possible biological controls such as T richoderma have
been shown to be antagonistic toward Armillaria, they are hard to establish in natural
situations because certain factors must interact in a relatively short time under a set of

specific environmental conditions (Munnecke 1981).

MOLECULAR METHODS USED FOR SPECIES IDENTIFCATION

Before the development of molecular techniques, Armillaria species were
distinguished using sexual compatibility (infertility) tests. These tests were described by
Buller (1931) and were first used to distinguish Armillaria species by Hintikka (1973),
and later expanded by Korhonen (1978). Serological studies have shown potential for
distinguishing Armillaria species as shown by Lung-Escarmant et al. (1978, 1985) and
Lung-Escarmant and Dunez (1979, 1980) by using an antiserum produced in rabbits to a
partially purified antigen of an A. mellea isolate. Using serology, Lung-Escarmant et al.
were able to distinguish between six European species of Armillaria (reviewed in
Burdsall et al.1990). Burdsall et al. (1990) used serology to distinguish between three
North American species of Armillaria using chicken eggs as a source of antibodies. In
recent years, many techniques have been developed for more rapid diagnosis of
Armillaria species targeting structural elements such as proteins, polysaccharides,

glycoproteins, and nucleic acids (Schulze and Bahnweg 1998). Methods have varied

25

from SDS-PAGE analysis of whole-cell proteins, isoenzyme pattern analysis, and the
restriction fragment patterns of mtDNA or nuclear rDNA.

Whole-cell protein SDS-polyacylamide electrophoresis pattern analysis was
attempted by Morrison et al. (1984), Poon (1988), and Lin et al. (1989), who found this
technique ineffective at distinguishing between Armillaria species. Lung-Escarmant et
al. (1985) used this technique with European species were only able to distinguish A.
gallica from other European species (reviewed in Schulze and Bahnweg 1998).

Isozyme analysis is another tool that can differentiate morphologically similar or
closely related species, varieties, and formae speciales. It can also be used to analyse
genetic variability, trace pathogen spread, follow the segregation of genetic loci, and
identify ploidy level of fungi and other pathogens (reviewed in Schulze and Bahnweg
1998). Harries and Hopkinson (1976) discuss how isoenzyme variation arises from three
phenomena: different alleles at a single locus (termed allozymes), multiple loci coding
for a single enzyme, and post-translational processing and formation of secondary
isoenzymes. Using isoenzyme analysis, six Armillaria intersterility groups in British
Columbia were distinguished using esterase and polyphenol oxidase by Morrison et al.
(1985). Lin et al. (1989) attempted to distinguish four NABS’s using isoenzyme analysis
of 20 different enzymes. Of the 20 enzymes tested, only alcohol dehydrogenase,
esterase, and polyphenol oxidase showed activity and only the esterase was able to
distinguish between species. Wahlstrdm et al. (1991) used pectin esterase and
polygalacturonase to differentiate between five European species (reviewed in Schulze

and Bahnweg 1998).

26

Anderson and Stasovski (1992) sequenced a portion of the Intergenic Spacer
(IGS) of the ribosomal RNA operon and suggested that restriction enzymes digests of this
region could be used to discriminate species. Harrington and Wingfield (1995) used the
primers developed by Anderson and Stasovski (1992) for the IGS-1 region and tested
their suggestion using restriction fragment length polymorphisms (RFLPs). The
restriction enzyme AluI produced patterns that could distinguish 6 of 11 Armillaria
species from European and American sources. Species not distinguishable with AluI
could usually be separated using other restriction enzymes (Harrington and Wingfield
1995). North American isolates of A. gallica and A. calvescens could not be
distinguished using this technique. Since Harrington and Wingfield (1995), other authors
have used this protocol to help characterize and distinguish species of Armillaria in North
America (Banik et al. 1996; Kim et al. 2000; Sierra et al. 1999; and White et al. 1998).
Banik et al. (1996) found that isolates of A. sinapina and A. gallica from the state of
Washington could not be distinguished from one another using banding patterns from
restriction digests of the IGS-1 region (Banik et al. 1996). White et al. (1998) were able
to use the IGS-2 region and restriction enzyme AluI to differentiate between A. sinapina
and A. gallica isolates with the same IGS-1 patterns. The use of restriction enzyme
digestion of the IGS region is currently the most used system to distinguish species of
Armillaria (White et al. 1998).

Sequencing of the Internal Transcribed Spacer (ITS) region has also been used to
show species differentiation. Chillali et al. (1998a,b) used the ITS region to differentiate
between European species and found that the ITS-2 region showed more variability

between the species than in previous work looking at the ITS-1 region. Lochman et al.

27

(2004) used PCR-RFLP of the ITS rRNA genes to analyze six Armillaria species from
the Czech Republic (Lochman et al. 2004). They found that restriction enzyme Hinfl was
able to discriminate between all six species. The attempt to design species-specific
primers for six European species was partially successful using variation within the IGS
and ITS regions of Armillaria isolates. However, due to high similarity between A.
astoyae and A. borealis and between A. cepistipes and A. gallica, these species pairs
could not be distinguished (Sicoli et al. 2003).

Molecular phylogeny has also been used to examine relatedness of Armillaria
species. Phylogenetics is an approach that allows one to recognize fungal species based
on concordance of multiple gene genealogies and compare them to other methods such as
morphology and reproductive behavior. The concept of species in fungi depends heavily
on the method that was employed to distinguish the species. Mayden (1997)
characterized different species concepts as either operational or theoretical. The only true
theoretical species concept was the Evolutionary Species Concept, which defines a
species as a single lineage of ancestor-descendent populations that maintain identity from
other lineages, thus having its own evolutionary tendencies and historical fate (Wiley
197 8). Other species concepts are considered secondary to the Evolutionary Species
Concept by Mayden (1997) which include the Morphological Species Concept (based on
variation in morphology), Biological Species Concept (based on mating capability), and
the Phylogenetic Species Concept (reviewed in Taylor et al. 2000). To date, phylogenetic
studies of Armillaria species agree with classifications based on the Biological Species
Concept. The most studied sequences for Armillaria include the ITS and IGS region

(Anderson and Stasovski 1992; Chillali et al. 1998; Coetzee et al. 2003, 2005ab; Hanna et

28

al. 2003, 2004; and Piercey-Normore et al. 1997). From these studies the authors were
able to infer a historical framework of species divergence. According to Hanna et al.
(2003), the ITS region along with another region [the nuclear large ribosomal subunit
(LSU)] were the most promising for evaluating evolutionary relationships among
Armillaria species, while the IGS-1 region was found to be more useful for revealing
intra-specific relations. Hanna et al. (2004) were also able to show genetic variability
among western A. ostoyae isolates from direct sequencing of these three regions with the
strongest variations in the ITS and IGS-1 regions. The next step is to examine other
regions and genes within the genome for variation between species. Maphosa et al.
(2006) has begun some of this work by studying the relationship of 42 isolates of
Armillaria, representing the majority of the species, using the elongation factor l-alpha
DNA sequence and found the results similar to previous comparisons using sequences of

the ITS and IGS-1 region (Maphosa et al. 2006).

SOMATIC INCOMPATIBILITY

Somatic incompatibility (S1) is referred to by Worrall (1997) as “the prevention of
effective fusion and integration following allorecognition (recognition of nonself)
between genetically distinct, conspecific tissues when isogenic (self) contacts result in
such fusion. ‘Somatic’ specifies a nonreproductive domain, distinguished the system
from sexual incompatibility.” This phenomenon is widespread among various organisms
including animals, slime molds, plants, and fungi. Somatic incompatibility in fungi can
be defined as the mycelial rejection between genetically distinct individuals within the

same species, and is used to maintain individuality of mated or secondary mycelium from

29

genetic exchange. In Armillaria, as well as other fungi, when two isolates of secondary
mycelium come in contact they are said to be incompatible if anastomose fail and a
barrage zone (zone of inhibition) forms between them. The isolates are said to be
compatible if the two mycelia merge and anastomoses persist between them. This
technique has been used among Armillaria and other fungi to identify clonal isolates
(reviewed in Worrall 1997).

Before molecular technology, the only other test, besides somatic incompatibility
reactions, for identifying genetic individuality were mating-type allele tests. Mating-type
allele tests involve four steps as described by Guillaumin et al. (1996): collection of
fruiting bodies, single spore isolation of all mating types, identification of the four mating
types from single spore isolates, and pairing of mating types with each other, involving
16 different haploid-haploid matings (Guillaumin et al. 1996). These two methods were
first used by Korhonen (1978) to look at clonal spread, and later others followed with
similar studies using either one or both of the methods (Anderson et. al. 1979b; Berthelay
and Guillaumin 1985; Guillaumin and Berthelay 1990; Kile 1983, 1986; Rizzo and
Harrington 1993; Thompson 1984; Ullrich and Anderson 1978). Smith et. al. (1990)
were among the first to use molecular markers to compare Armillaria genets (clones).
They compared mating-type allele tests to mitochondrial DNA restriction fragment
patterns and found perfect correspondence between the two methods, despite the fact that
mating types are encoded by nuclear DNA. Later studies found that correspondence
between these two methods is not always the case and that the number of mtDNA types
often tends to be lower than the number of genets found by the previous study

(Guillaumin et. al. 1996; Smith et al. 1994). Other studies found that using DNA

30

fingerprinting or analysis of nuclear DNA using random amplified polymorphic DNA
(RADP) analysis and microsatellites tend to correspond much better to somatic
incompatibility reactions and mating-type allele tests.

Today the fastest and most convenient method, if available, to distinguish
between SI groups is through the use of microsatellites, also known as simple sequence
repeats (SSRs). These are short repeats of DNA nucleotides usually 20-60 bp in length
with tandem repetition of 1-5 base pairs such as (GT)n, (CA)n, (CAA)n, or (GACA)n.
These repeats are a common feature in all eukaryotes occurring in as many as 105
different locations. However, basidiomycetes rarely contain more than 5% repetitive
DNA, unlike the older division Zygomycota which usually contains 30% or more
repetitive DNA. Regions with highly repetitive DNA have a high frequency of mutation
allowing these regions to be used to differentiate between taxa or an individual within a
single species (reviewed in sttemeyer and Kreibich 2002). In studying 19 isolates of
A. ostoyae belonging to six SI groups, Langrell et al. (2001) observed 12 distinct loci
harboring a repetitive motif and were able to develop a set of 12 primers to distinguish SI
groups. Langrell et. al. (2001) also observed that with the 12 primers they developed
there Was a high level of cross-species amplification with closely related Armillaria
species, and suggests this set of primers may be able to explore SI groups across the
genus. Lefrancois et al. (2002) later explored this idea with A. gallica using six of the
original primers developed for A. ostoyae, and were able to improve a set of five novel
primer pairs for A. gallica. Worrall et al. (2004) used two of the di—nucleotide primer sets
developed by Langrell (2001) and also developed two primer sets for tri-nucleotide

repeats to distinguish clones of A. ostoyae in a Colorado campground. Although SSRs

31

are a fast and easy way to identify individual genets, at this time they still should be
backed up with previous methods. With more exploration within species and also across

the genus the use of SSRs can be better utilized.

DISTRIBUTION STUDIES

There were few early distribution studies which examined the spread of
Armillaria mellea senso lato in orchards. In 1925, Hendrickson published a paper on the
spread of Armillaria in a plum and apricot orchard in California over a period of 23
years. Nearly 66% of the trees died over the course of the study. Due to multiple sources
of infection, Hendrickson was unable to determine a linear rate of spread of an individual
clone (reviewed in Marsh 1952). In 1952, Marsh published a study of the spread of
Armillaria in apple (Malus spp.) orchards and in a black currant (Ribes nigrum L.)
planting, concluding that the pattern of spread was primarily due to root to root contact.
He also found that the trees killed in the orchard tended to be over 15 years old, half of
them were over 30 years old. Marsh also found that the rate of spread tended to be higher
in black currants than apples at about 1.8 m per year (Marsh 1952). Rishbeth (1968)
observed the rate of spread from various locations in England to be 1.1 m to 1.6 m per
year in ash, Norway spruce (Picea abies (L.) Karst.), and Douglas-fir (reviewed in Kable
1974). Kable (1974) studied the natural spread of Armillaria (Armillariella) from a
single source in an irrigated peach orchard in Australia. This study suggested that peach
is highly susceptible and that the age of the tree did not influence its susceptibility.
Spread was found to be at a rate of 0.8 m to 3.2 m per year from grth of rhizomorphs

and was found to be more rapid in soil receiving direct furrow irrigation (Kable 1974).

32

The radial rate of spread for A. ostoyae has been estimated by Shaw and Roth (1976) and
Feet et al. (1996) at approximately 1 m per year in young conifer stands and 0.22 m per
year in a 110 year old Douglas-fir stand.

After Armillaria mellea senso Iato was divided into a number of new biological
species, many distribution surveys were published. Guillarnin et al. (1993) compiled
over 4000 records of the six European species of Armillaria to develop distribution maps.
Qualities such as geographical and altitudinal distribution, host range, pathogenicity,
dissemination, and ecological roles were also compiled and discussed (Guillamin et al.
1993). In North America, species distribution studies have mostly been conducted in
forested areas (Blogdett and Worrall 1992; Harrington and Rizzo 1993; McLaughlin
2001). In New York, Blodgett and Worrall (1992) collected samples of Armillaria from
273 forested sites, and examined the host and species relationship for six North American
biological species of Armillaria. Similar studies conducted by Harrington and Rizzo
(1993) identified six species of Armillaria from the state of New Hampshire, and in
Southern Ontario McLaughlin identified six species of Armillaria. The host range in all
three studies found that A. gemina, A. calvescens, A. mellea, A. gallica, and A. sinapina
were alrnost always found on hardwood species, while A. ostoyae was the only species
predominantly found on conifers. Proffer et al. (1987) assessed the distribution of
Armillaria species in Michigan tart cherry orchards. In this study, three species of
Armillaria were found to be vigorous pathogens to orchard grown Prunus, including A.
ostoyae, A. mellea, and A. calvescens. Although A. ostoyae is a principal host to conifer,

Proffer et al. (1987) found that it is the predominant pathogen to tart cherry in Leelanau,

33

Grand Traverse, and Benzie counties where cherry production is greatest and the
pathogen is widespread.

Currently many studies are looking at the spread and distribution of species of
Armillaria within distinct areas. These studies have focused largely on clonal spread in
forests and some within orchards (Anderson et al. 1979b; Bendel 2006; Ferguson et al.
2003; Klein-Gebbinck et al. 1991; Lung-Escarmant and Gayon 2004; Prospero et al.
2003; Rizzo et al. 1998; Shaw and Roth 1976, Smith et al.1992; Smith et al. 1994;
Worrall 2004). Most of these studies focus on the clonal spread of A. 0st0yae in
coniferous forests or plantations. Klein-Gebbinck et al. (1991) determined the spread of
A. ostoyae clones in juvenile lodgepole pine (Pinus contorta Dougl. ex Loud.) in Alberta
Canada. From the two study sites they found nine genets at one site and five genets at
another site. The infected trees were aggregated, which was determined using a variance
mean ratio of 2.72. They also found that many of the discontinuous patches of trees were
infected by a single genet of Armillaria. Another study in northern Michigan, examined
the clonal spread of A. ostoyae in infected red pine (Pinus resinosa Aiton) seedlings
(Smith et al. 1994). The plantation was planted within a 1.2-ha clearing in a hardwood
forest. From this study 22 genets of A. ostoyae were found within the plantation and '
around the plantation vicinity. Smith et al. (1994) concluded that based on the estimates
of size and shape, that some of the genets were established at the time of stand
conversion, and that the largest clone may predate the existing hardwood forest, which
replaced a forest of mostly pine. Rizzo et al. (1998) examined the clonal spread of A.
mellea in two pear (Pyrus spp.) orchards, previously hardwood forests, in California. In

this study, Rizzo et al. (1998) discovered four genets at one site with three of the genets

34

being over 100 m in length. At the second site five genets were found, however these
genets were smaller, ranging in sizes from 20 to 60 min length. The conclusions of this
study suggest that based on the size of many of these genets, they were established prior
to planting of the orchards, however the size indicates that expansion has occurred since
the pears were planted. Another study examined the delineation and biology of genets of
three A. ostoyae, A. gemina, and A. calvescens at a mixed conifer-hardwood site and at a
spruce-fir site in New Hampshire (Rizzo and Harrington 1993). The results from this
study found six genets of A. ostoyae at the spruce-fir site, colonizing 34% of the conifers
and 13% of the hardwoods. At the mixed conifer-hardwood site A. ostoyae and A.
calvescens were found colonizing both conifers and hardwood hosts, while A. gemina
was only found colonizing only hardwood hosts. Six A. ostoyae genets, two A.
calvescens, and two A. gemina genets were identified using somatic incompatibility tests
and isoenzyme analysis. A study, which examined the clonal spread of A. gallica (A.
bulbosa) in the Upper Peninsula of Michigan, found that Armillaria was among the
world’s largest and oldest living organisms. This study found a clone or genet of A.
gallica to be a minimum of 15 hectares with a growth rate at approximately 0.2 m per
year and that had remained genetically stable for over 1,500 years (Smith et al. 1992).
The finding from this publication by Smith et al. (1992) was picked up by the popular
press and Armillaria was referred to as the “humongous fungus” (V olk 2003). Since then
a genet of A. astoyae has been found to extend over 37 hectares and was estimated to be
1000 to 2000 years old (Bendel 2006) and in Oregon a genet of A. ostoyae was found to

be up 965 hectares (Ferguson et al. 2003).

35

THE HOST RANGE

Armillaria has a broad host range both as a saprophyte and as a pathogen.
Armillaria species have been reported on conifer and hardwood hosts through out North
America, Europe, Asia, South America, Australiasia and Afiica (Hood et a1. 1991). In
North America, Armillaria is primarily known to act as a virulent pathogen on conifers in
the west, and as an opportunistic pathogen of already stressed hardwood species in the
east (Shaw and Roth 1978). Studies on Armillaria root rot in the United States have
focused on hardwood species, conifer species, and on fruit trees. In fruit crops grown in
the United States, Armillaria spp. have been reported on stone fruits (Prunus spp.), pome
fruits (Malus spp. and Pyrus spp.), kiwifruit (Actinidia spp.), avocado (Persea spp.),
citrus fruits (Citrus spp.), berryfruits (Ribes spp. and Rubus spp.), fig (Ficus carica),
guava (Psidium spp.), and strawberry (Fragaria spp.) (Hood et al. 1991). Within the
state of Michigan, saprophytic species and pathogenic species of Armillaria have been
reported on a variety of forest hosts and crops (Table 1-2).

Cherries, one of the major crops of Michigan agriculture, are severely impacted
by Armillaria root rot. Michigan leads the nation in tart cherry production, producing
about 80 % of the nation’s tart cherries, with 30,800 acres devoted to tart cherry
production. In 2001 , 297 million pounds of tart cherries were produced, with a total
value of production at approximately $44 million (Crop Profile for Tart Cherries in
Michigan 2003). Michigan also produces approximately 17% of the nation’s sweet
cherries (7,400 acres). Leelanau County in Michigan contains approximately 51% of

Northwest Michigan’s cherry acreage. Michigan has both favorable soil and climatic

36

Table 1-2: Reported host range of both saprophytic and pathogenic species of

Armillaria in Michigan: Modified by Kromroy (2004)

 

Species

NABSI

Host

Location Found

Reference

 

A. calvescens

lll

Prunus cerasus

Lower Peninsula

Proffer et al. 1987

 

Acer saccharum
Betula papyrifera
Lonicera spp.
Populus tremuloides
Quercus rubra
Quercus veluntina
Tilia Americana
Ulmus spp.

Upper Peninsula

Banik et al. 1995

 

A. gallica

VII

Pinus resinosa

Lower Peninsula,
Northern
Michigan

Smith et al. 1992

 

Abies balsamea
Pinus resinosa
Pinus strobus

Acer rubrum
Acer saccharinum
A cer saccharum
Betula papyrrfera
C arya ovata
F raxinus
pennsylvanica
Populus tremuloides

Prunus serotina

Quercus alba
Quercus ellipor'dalis
Quercus macrocarpa

Quercus rubra
Quercus veluntina
Ulmus spp.

Upper Peninsula

Banik et al. 1995

 

A. mellea

VI

Prunus cerasus
Quercus spp.

Lower Peninsula

Proffer et al. 1987

 

Acer saccharum
Quercus spp.

Upper Peninsula

Banik et al. 1995

 

A. ostoyae

 

 

Prunus cerasus
Prunus avium

Prunus persica
Malus pumila

Lower Peninsula

Proffer et al. 1987

 

 

Pinus resinosa

 

Lower Peninsula,
Northern
Michigan

 

Smith et al. 1990

 

37

 

Continuation of Table 1-2: Reported host range of both sapr0phytic and pathogenic

species of Armillaria in Michigan: Modified by Kromroy (2004)

 

Species

NABS'

Host

Location Found

Reference

 

A. ostoyae

Acer rubrum
Betula papyrifera
Pinus resinosa
Populus sp.

Lower Peninsula,
Northern
Michigna

Smith et al. 1994

 

Abies balsamea
Acer rubrum
A cer saccharum
Betula allegham'ensis
Betula papyrifera
F raxinus nigra
Picea banksiana
Picea glauca
Pinus resinosa
Pinus strobus
Populus tremuloides
Quercus rubra
Quercus veluntina
Thuja occidentalis
Tsuga C anadensis
Ulmus spp.

Upper Peninsula

Banik et. al. 1995

 

 

A. sinapina

 

Populus sp.

Lower Peninsula,
Northern
Michigan

Smith et al. 1994

 

 

Abies balsamea
Thuja occidentalis
Tsuga C anadensis

Acer rubrum
Betula allegham'ensis
F raxinus nigra
Populus tremuloides

 

Quercus rubra

Upper Peninsula

 

Banik et al. 1995

 

1. NABS = North American Biological Species. See Table 1 for more information on NABS.

38

 

conditions for growing cherries. Lake Michigan provides moderate temperatures with
frost-free autumns and a delayed bloom period in the spring (Cherries 2006).

Cherries, along with other stone fruits, belong to the genus Prunus in the family
Rosaceae (Zomlefer 1994). There are two species of cultivated cherry: the sweet cherry,
Prunus avium L., and the sour cherry, Prunus cerasus L. Both species originated
fromEurope and Southwest Asia. In the United States, the tart cherry industry is virtually
a monoculture with ‘Montmorency’ being the dominant variety grown. However in
recent years Morello varieties are also being grown in Michigan. The variety
‘Montmorency’ is a 400 year old variety from France (Iezzoni 2006).

All commercially grown cherries are grafted. A rootstock makes up the healthy
root system to which the upper part of the tree, or scion, is grafted. The rootstock and
scion can be different species but must be closely related to be compatible. Rootstocks
influence many aspects of the scion’s growth and vigor, including productivity, precocity,
tree size, tree architecture, fruit size, and fruit quality, and also horticultural decisions
such as pruning, training, tree support, and labor management. Cherry rootstocks in
Michigan have been selected for precocity, productivity, vigor control, disease tolerance,
and adaptability to different soils or climates (Lang 2007). In Michigan, P. mahaleb L.
(mahaleb) rootstock is the primary rootstock used with tart cherry. Mahaleb rootstock
does well in the sandy and acidic soil conditions in which tart cherry trees are usually
grown and is also known to keep the trees smaller and easier to manage. Sweet cherry in
Michigan is usually grafted onto P. avium L. (mazzard) rootstock.

Cherry rootstocks play a role in susceptibility to Armillaria root rot. Proffer et al.

(1988) tested a variety of Prunus rootstocks including mahaleb, mazzard, and 17 hybrids.

39

In that inoculation study mahaleb rootstock was more susceptible than mazzard rootstock.
Mazzard is however found to be infected with Armillaria in the field (Nugent 2005). The
tested hybrids were found to be neither immune nor highly resistant to infection (Proffer
et al. 1988).

The possible susceptibility of grapevine (Vitis spp.) to Armillaria root rot is a new
concern of Michigan growers. Michigan is the fourth largest grape producer in the
United States, with approximately 14,400 bearing acres. In 2005 over 102,000 tons of
grapes were produced. Michigan produces mostly juice grapes, with ‘Concord’ and
‘Niagara’ the dominant grape varieties. Although Michigan mostly produces juice
grapes, its production of wine grapes is growing, especially in the cherry regions where
Armillaria is often a problem (Creighton 2007). Leelanau County has 275 acres devoted
to grapes (MDA-Grape & Wine 2007) and the Leelanau Peninsula and the Old Mission
Peninsula produce 51% of Michigan’s wine grapes, while another 45% of wine grapes
are grown along the southwest Michigan shoreline and in Fennville (Michigan Wineries
2007). Because of the growing wine industry in Michigan, Armillaria may soon be found
to be a problem for many Michigan growers who are converting cherry orchards into
vineyards.

Although this may be a new concern to Michigan growers, Armillaria root rot has
been reported on grape in Australia, Brazil, Central and Eastern Europe, Southeast
California, and the West coast of California of the United States. In 1880, Millardet first
reported a root rot disease on samples of Vitis species from Belgium, Bulgaria, England,
France, Germany, Greece, Hungary, Italy, Scotland, Spain, and Switzerland. The

causative disease described by Millardet was later identified as A. mellea (reviewed in

40

Agur'n—Casal 2004). More recently, Aguin-Casal et al. (2004) reported A. gallica and A.
cepistipes for the first time on Vitis spp. in Spain. In California, Baumgartner et al.
(2006ab) have begun to test grape rootstock susceptibility to A. mellea. Baumgartner
tested eight different rootstocks finding rootstocks 3309C and Riparia Gloire, commonly
used in Michigan, most susceptible to A. mellea, while Freedom (not commonly used in
Michigan) was most resistant. Other rootstocks that commonly do well in Michigan soils,
that were not tested by Baumgartner, include 5-BB, 804, 101-14 Mgt ( Howell et al.
1999). Eventually these rootstocks as well as the rootstocks tested by Baumgartner

should be tested for susceptibility to Michigan isolates of Armillaria.

41

REFERENCES

Adaskaveg, J.E., Forster, H., Wade, L., Thompson, D.F., & Connell, J .H. (1999).
Efficacy of sodium tetrathiocarbonate and propiconazole in managing Armillaria
root rot of almond and peach rootstock. Plant Disease. 83:240-246.

Aguin, 0., Mansilla, J .P., & Sainz, M.J. (2006). In vitro selection of an effective

fimgicide against Armillaria mellea and control of white root rot of grapevine in
the field. Pest Management Science. 62: 223-228.

Aguin-Casal, 0., Sainz-Oses, M.J., & Mansilla—Vazquez, J .P. (2004). Armillaria species
infesting vineyards in northwestern Spain. European Journal of Plant Pathology.
110: 683-687.

Anderson, J. B. & Stasovski, E. (1992). Molecular Phylogeny of Northern Hemisphere
species of Armillaria. Mycologia. 84:505-516.

Anderson, J .B. & Ullrich, RC. (1979) Biological species of Armillaria mellea in North
America. Mycologia. 71 : 402-414.

Anderson, J .B., Ullrich, R.C., Roth, L.F., & Filip, G.M., (1979b). Genetic identification
of clones of Armillaria mellea in coniferous forests in Washington.
Phytopathology. 69: 1 109-1 11 1.

Anderson, J .B. & Ullrich, RC. (1982). Diploids of Armillaria mellea: synthesis, stability,
and mating behavior. Canadian. Journal of Botany. 60: 432-439.

Banik, M.T., Paul, J .A., & Burdsall, H.H. Jr. (1995). Identification of Armillaria species
from Wisconsin and adjacent areas. Mycologia. 87: 707-712.

Banik, M.T., Volk T.J., & Burdsall, H.H. Jr. (1996). Armillaria species of the Olympic
Peninsula of Washington state, including confirmation of North American
biological species XI. Mycologia. 88: 492-496.

Baumgartner, K. (2004). Root collar excavation for postinfection control of Armillaria
root disease of grapevine. Plant Disease. 88: 1235-1240.

Baumgartner, K. & Rizzo, D.M. (2006a). Relative resistance of grape vine stock to
Armillaria root disease. American Journal of Enology and Viticulture. 7: 408-414.

Baumgartner K. & Rizzo, D. M. (2006b). Resistant grapevine rootstocks for control of

Armillaria root disease. American Journal of Enology and Viticulture. 57:
383A-3 83A.

42

Bendel, M., Kienast, F. & Rigling, D. (2006). Genetic population structure of three
Armillaria species at the landscape scale: a case study from Swiss Pinus mugo
forests. Mycologial Research. 110: 705-712.

Berthelay, S., & Guillaumin, J .J . (1985). Geographic-distribution of the incompatibility
factors of the diploid Basidiomycete Armillaria obscura (Secretan) Herink.
Cryptogamie Mycologie. 6: 185-196.

Bliss, D. E. (1951). The Destruction of Armillaria mellea in Citrus Soils. Phytopathology.
41: 665-683.

Blodgett, J. T. & Worrall, J .J . (1992). Distributions and Hosts of Armillaria species in
New York. Plant Disease. 76: 166-170.

Buller, A. H. R. (1931), Researches onfungi., Vol. 4, London: Longrnans Green.

Burdsall, H.H. Jr., Banik, M., & Cook, ME. (1990). Serological differentiation of three
species of Armillaria and Lentinula edodes by enzyme-linked immunosorbent

assay using immunized chickens as a source of antibodies. Mycologia. 82: 415-
423.

Callac, P., Spataro, C., Caille, A., & Imbemon, M. (2006). Evidence for outcrossing via
the Buller Phenomenon in a substrate simultaneously inoculated with spores and

mycelium ongaricus bisporus. Applied and Environmental Microbiology. 72:
2366-23 72.

Cherries. (July 2006). Retrieved July 3, 2007 from the Leelanau Government and
Community Center webpage:
http://www.leelanaucounty.com/agriculture001 5 .asp.

Chillali, M., Idder-Ighili, H., Guillaumin, J .J ., Mohammed, C., Lung-Escarrnant, & B.,
Botton, B. (1998a). Variation in the ITS and IGS regions of ribosomal DNA

among the biological species of European Armillaria. Mycological Research. 102:
533-540.

Chillali, M., Wipf, D., Guillaumin, J .J ., Mohammed, C., & Botton, B. (1998b).
Delineation of the European Armillaria species based on the sequences of the
internal transcribed spacer (ITS) of ribosomal DNA. New Phytologist. 138: 553-
561.

Coetzee, M. P. A., Wingfield, B.D., Bloomer, P., Ridley, G.S., & Wingfield, M.J. (2003).

Molecular identification and phylogeny of Armillaria isolates from South Africa
and lndo-Malaysia. Mycologia. 95: 285-293.

43

Coetzee, M. P. A., Wingfield, B.D., Bloomer, P., & Wingfield, M.J. (2005a).
Phylogenetic analyses of DNA sequences reveal species partitions amongst
isolates of Armillaria from Afiica. Mycological Research. 109: 1223-1234.

Coetzee, M. P. A., Wingfield, B.D., Kirists, T., Chhetri, D.B., Bloomer, P., & Wingfield,
M.J. (2005b). Identification of Armillaria isolates from Bhutan based on DNA
sequence comparisons. Plant Pathology. 54: 34-45.

Creighton, D. (March 2007). National study: Michigan grape and wine industry
contributes $789 million and 5,400 jobs. Retrieved July 3, 2007 from the
Michigan Department of Agriculture webpage:
http://michigan.gov/mda/0,l607,7-125-1527_3628-163468--,00.html.

Crop profile for tart cherries in Michigan. (January 2003). Retrieved July 3, 2007 from
the National Information System for the Regional IPM Centers webpage:
http://www.ipmcenters.org/cropprofiles/docs/MITartCherry.html.

Ferguson, B.A., Dreisbach, T.A., Parks, C.G., Filip, G.M., & Schmitt, CL. (2003).
Coarse-scale population structure of pathogenic Armillaria species in a mixed-
conifer forest in the Blue Mountains of northeast Oregon. Canadian Journal of
Forest Research. 33: 612—623.

Filip, G. M. & Roth, LR (1977). Stump injections with soil fumigants to eradicate
Armillariella mellea from young-growth ponderosa pine killed by root rot.
Canadian Journal of Forest Research. 7: 226-231.

Garraway, M.O., Hiitterman, A., & Wargo, RM. (1991). Ontogeny and physiology. In:
Armillaria Root Disease. Washington, DC, United States Department of
Agriculture Forest Service Agriculture Handbook No. 691. 21-46.

Garrett, S. D. (1956). Rhizomorph behavior in Armillaria mellea (V ahl) Que]: Logisitcs
of infection. Annals of Botany, NS. 20: 193-209.

Gregory, S.C., Rishbeth, J. & Shaw, C.G. III. 1991. Pathogenicity and virulence. In:
Armillaria Root Disease. Washington, DC, United States Department of
Agriculture Forest Service Agriculture Handbook No. 691. 76-87.

Grillo, R., Korhonen, K., Hantula, J ., & Hietala, A.M. (2000). Genetic evidence for
somatic haploidization in developing fruit bodies of Armillaria tabescens. Fungal
Genetics and Biology. 30: 135-145.

Gubler, W.D. (1992). Armillaria Root Rot. In Grape Pest Management. D.L. F laherty,
L.P. Christensen, W.T. Lanini, J .J . Marois, P.A. Phillips and LT. Wilson (Eds).
pp. 92-93. University of California, Division of Agriculture and Natural
Resources, Publication #3343, Oakland.

44

Guillaumin, J .-J ., Anderson, J .B., Legrand, P., Ghahari, S., & Berthelay, S. (1996). A
comparison of different methods for the identification of genets of Armillaria spp.
New Phytologist. 133: 333-343.

Guillaumin, J .J ., & Berthelay, S. (1990). Comparison of 2 methods for the identification
of the clones of the parasitic basidiomycete Armillaria obscura (=A-ostoyae).
European Journal of Forest Pathology. 20: 257-268.

Guillaumin, J .-J ., C. Mohammed, C, Anselmi, N., Courtecuisse, R., Gregory, S.C.,
Holdenridder, 0., Intini, M., Lung, B., Marxmuller, H., Morrison, D., Rishbeth,
J ., Terrnorshuizen, A.J., Tirro, A, & Van Dam, B. (1993). Geographical
distribution and ecology of the Armillaria species in western Europe. European
Journal of Forest Pathology. 23: 321-341.

Hanna, J .W., Kim, M.-S., Klopfenstein, N.B., Zambino, P.J., & McDonald, G.I. (2003).
Characterization of North American Armillaria species: Phylogenetic
relationships from IGS, ITS, and nuclear large subunit ribosomal DNA sequences.
Proceedings of the annual meeting of the American Phytopathological Society. 9-
13 August 2003, Charlotte, NC. Phytopathology. 93: S33.

Hanna, J .W., Klopfenstein, N.B., Kim, M.-S., McDonald, G.I., & Moore, J .A. (2004).
Phylogeography of Armillaria astoyae in the western United States. Annual
meeting of the American Phytopathological Society, 31 July to 4 August 2004,
Anaheim, CA. poster paper. Phytopathology. 94:838.

Hagle, S.K., & Shaw, C.G. III. (1991). Avoidance and reducing losses from Armillaria
root disease. In: Armillaria Root Disease. Washington, DC, United States
Department of Agriculture Forest Service Agriculture Handbook No. 691 . 157-
172.

Harrington, T. C. & Rizzo, D.M. (1993). Identification of Armillaria species from New
Hamshire. Mycologia. 85: 365-368.

Harrington, T. C. & Wingfield, 8D. (1995). A PCR-based identifcation method for
species of Armillaria. Mycologia. 87: 280-288.

Hintikka, V. (1973). A note of the polarity of Armillariella mellea. Karstenia. 13: 32-39.

Hood, I.A., Redfem, D.B., & Kile, GA. (1991). Avoidance and reducing losses from
Armillaria root disease. In: Armillaria Root Disease. Washington, DC, United
States Department of Agriculture Forest Service Agriculture Handbook No. 691.
1 5 7-172.

Howell, G.S., Miller, D.P., & Zabadal, T.J. (November 1999). Wine grape varieties for

Michigan. Retrieved July 3, 2006 from the Michigan State University Extension
webpage: http://webl .msue.edu/imp/modfr/26439701 .html.

45

Iezzoni, AF. (2007). Amy Iezzoni. Retrieved July 3, 2007 from the Department of
Horticulture at Michigan State University webpage:
http://www.hrt.mus.edu/faculty/OLD/main__lezzoni.htm.

1PM: Landscape and Turf: Armillaria Root Rot of Trees and Shrubs. (March 2000)
Retrieved November, 3, 2005, from Integrated pest management at the University
of Illinois Urbana-Charnpaign Website:
http://www.imp.uiuc.edu/landturf/disease/armillaria/

Jones, AL. & Sutton, T.B. (1996). Diseases of Tree Fruits in the East. East Lansing, MI,
Michigan State Univeristy.

Kable, P. F. (1974). Spread of Armillariella sp. in a peach orchard. Transactions of the
British Mycological Society. 62: 89-98.

Kile, GA. (1983). Identification of genotypes and clonal development of Armillaria
luteobubalina Watling & Kile in eucalypt forest. Australiasian Journal of Botany.
3 1 : 65 7-671 .

Kile, GA. (1986). Genotypes of Armillaria hinnulea in wet sclerophyll eucalyptis forest
. in Tasmania. Transactions of the British Mycological Society. 87: 312-314.

Kile, G.A., McDonald, G.I. & Byler, J .W. (1991). Ecology and disease in natural forest.
In: Armillaria Root Disease. Washington, DC, United States Department of
Agriculture Forest Service Agriculture Handbook No. 691.102-121.

Kim, M.-S., Klopfenstein, N.B., McDonald, G.I., Arumuganathan, K., & Vidaver, AK.
(2000). Characterization of North American Armillaria species by nuclear DNA
content and RF LP analysis. Mycologia. 92: 874-883.

Klein-Gebbinck, H. W., Blenis, P.V., & Hiratsuka, Y. (1991). Clones of Armillaria
ostoyae and the pattern of infected juvenile lodgepole pine in Alberta, Canada.
European Journal of Forest Pathology. 21: 260-266.

Korhonen, K. (1978). Interfertility and clonal size in the Armillaria mellea complex.
Karstenia. 18: 31-42.

Korhonen, K. (1980). The origin of clamped and clampless basidia in Armillariella
0stoyae. Karstenia. 20: 23-27.

Kromroy, K. W. (2004). Identification of Armillaria species in the Chequamegon-Nicolet

National Forest. Res. Note NC-388. St. Paul, MN: US. Department of
Agriculture, Forest Service, North Central Research Station. 10 p.

46

Lang, GA. (2007). Sweet Cherry Rootstock. Retrieved July 3, 2007 from Dr. Greg Lang
at the Department of Horticulture at Michigan State University webpage:
http://www.hrt.mus.edu/faculty/langg/Sweet_Cherry_Rootstocks.html.

Langrell, S. R. H., Lung-Escarrnant, B., & Decroocq, S. (2001). Isolation and
characterization of polymorphic simple sequence repeat loci in Armillaria
ostoyae. Molecular Ecology Notes. 1: 305-307.

Lefrancois, J .—B., Lung-Escarrnant, B., & Langrell, S.R.H. (2002). Development of novel
polymerase chain reaction (PCR) based microsatellite markers in Armillaria

gallica by cross-species amplification and species-specific cloning. Molecular
Ecology Notes. 2: 182-184.

Lochman, J ., Sery, O., Jankovsky, L., & Mikes, V. (2004). Variations in rDNA of Czech
Armillaria species determined by PCR and HPLC. Mycological Research. 108:
1153-1161.

Lung-Escarrnant, B. & D. Guyon, D. (2004). Temporal and Spatial Dynamics of Primary
and Secondary Infection by Armillaria ostoyae in a Pinus pinaster Plantation.
Phytopathology. 94: 125-131.

Maphosa, L., Wingfield, B. D., Coetzee, M. P. A., Mwenje, E., & Wingfield, M. J.
(2006). Phylogenetic relationships among Armillaria species inferred from partial

elongation factor l-alpha DNA sequence data. Australasian Plant Pathology. 35:
513-520.

Marsh, R. W. (1952). Field observations on the spread of Armillaria mellea in apple
orchards and in a blackcurrant plantation. Transactions of the British Mycological
Society. 201 -207.

McLaughlin, J. A. (2001). Distribution, hosts, and site relationships of Armillaria spp. in

central and southern Ontario. Canadian Journal of Forest Research. 31: 1481-
1490.

MDA-Grapes & Wine. (2007). Retrieved on July 3, 2007 from the Michigan Department
of Agriculture webpage: http://www.michigan.gov/mda/O,l607,7-125-
1570_2468_2469-12707--,00.html.

Michigan Wineries. (2007). Retrieved July 3, 2007 from the Absolute Michigan
webpage: http://www.absolutemichigan.com/search/?articleid=2888.

Morrison, D. J ., (1989). Pathogenicity of Armillaria species is related to rhizomorph
growth habit. In: Proc. 7th Int. Conf. Root and Butt Rots, Victoria and Vernon,
British Columbia, Canada, 9—16. August 1988. Ed. by Morrison, D. J. Victoria,
BC: Forestry Canada, pp. 584—589.

47

Morrison, DJ. (2004). Rhizomorph grth habit, saprophytic ability and virulence of 15
Armillaria species. Forest Pathology. 34: 15-26.

Morrison, D.J., Williams, R.E. & Whitney, RD. (1991). Infection, disease development,
diagnosis, and detection. In: Armillaria Root Disease. Washington, DC, United
States Department of Agriculture Forest Service Agriculture Handbook No. 691.
62-75.

Munnecke, D. E., Kolbezen, M.J., Wilbur, W.D. & 0hr, H.D. (1981). Interactions
involved in controlling Armillaria mellea. Plant Disease. 65: 384-3 89.

Munnecke, D. E., Kolbezen, M.J., & Wilbur, W.D. (1973). Effect of methyl bromide or
carbon disulfide on Armillaria and T richoderma growing on agar medium and

relation to survival of Armillaria in soil following fiunigation. Phytopathology.
63: 1352-1357.

Munnecke, D. E., Wilbur, W., & Darley, HF. (1976). Effect of heating or drying on
Armillaria mellea or T richoderma viride and the relation to survival of A. mellea
in soil. Phytopathology. 66: 1363-1368.

Peabody, D. C., Motta, J. J ., & Therrien, C. D. (1978). Cytophotometric evidence for
heteroploidy in the life cycle of Armillaria mellea. Mycologia. 70: 487-498.

Peabody, D. C., & Peabody, R. B. (1984). Microspectrophotometric nuclear cycle
analyses of Armillaria mellea. Experimental Mycology. 8: 161-169.

Peabody, D.C., & Peabody, R. B. (1985). Widespread haploidy in monokaryotic cells of
mature basidiocarps of Armillaria bulbosa, a member of the Armillaria mellea
complex. Experimental Mycology. 9: 212-220.

Peabody, D.C., & Peabody, RB. (1986). Quantitative differences in nuclear DNA
content of three stages in the life cycle of Armillaria bulbosa. Mycologia. 78:
967-968.

Peabody, R. B., & Peabody, D. C. (1987). Haploid monokaryotic basidiocarp tissues in
species of Armillaria. Canadian Journal of Botany. 65: 69-71.

Peabody, R. B., & Peabody, D. C. (2003). Research interests. Retrieved July 26, 2007
from the Stonehill College Biology Department webpage:

httpl/faculty.stonehill.edu/dpeabody/research_long.htrn.

Peabody, R. B., Peabody, D.C. & Sicard KM. (2000). A Genetic Mosaic in the Fruiting
Stage of Armillaria gallica. Fungal Genetics and Biology. 29: 72-80.

48

Peet, F.G., Morrison, D.J., & Pellow, K.W. (1996). Rate of spread of Armillaria ostoyae
in two Douglas-fir plantations in the southern interior of British Columbia.
Canadian Joru'nal of Forest Research. 26: 148-151.

Piercey-Norrnore, M. D., Egger, K.N., & Berube, J .A. (1998). Molecular Phylogeny and
Evolutionary Divergence of North American Biological Species of Armillaria.
Molecular Phylogenetics and Evolution. 10: 49-66.

Proffer, T. J ., Jones, A.L., & Ehret, GR. (1987). Biological species of Armillaria isolated
from sour cherry orchards in Michigan. Phytopathology. 77: 941-943.

Proffer, T. J ., Jones, A.L., & Perry, R.L. (1988). Testing of cherry rootstocks for
resistance to infection by species of Armillaria. Plant Disease. 72: 488-490.

Prospero, S., Rigling, D., & Holdenridder, O. (2003). Population structure of Armillaria
species in managed Norway spruce stands in the Alps. New Phytologist. 158:
365-373.

Redfem, D. B. (1971). Chemical Control of Honey Fungus (Armillaria mellea).
Procedings from the 6th Br. Insecticide and Fungicide Conference. 469-474.

Redfem, D.B., & Filip, GM. (1991) Inoculum and infection. In: Armillaria Root Disease.
Washington, DC, United States Department of Agriculture Forest Service
Agriculture Handbook No. 691. 48-60.

Rishbeth, J. (1970). The role of basidiospores in stump infection by Armillaria mellea. In:
Toussoun, T.A.; Bega, R.V.; Nelson, P.E. eds. Root diseases and soil-bome
Pathogens: Proceedings of the symposium; 1968 July; London: Imperial College.
Berkeley: University of California Press: 141-146.

Rishbeth, J. (1976). Chemical treatment and inoculation of hardwood stumps for control
of Armillaria mellea. Annals of Applied Biology. 82: 57-70.

Rishbeth, J. (1985). Infection cycle of Armillaria and host response. European Journal of
Forest Pathology. 15: 332-341.

Rizzo, D. M. & Harrington, TC. (1992). Nuclear migration in diploid-haploid pairings of
Armillaria ostoyae. Mycologia. 84: 863-868.

Rizzo, D. M. & Harrington, TC. (1993). Delineation and biology of clones of Armillaria
ostoyae, A. gemina, and A. calvescens. Mycologia. 85: 164-174.

Rizzo, D. M. & May, G. (1994). Nuclear replacement during mating in Armillaria
ostoyae (Basidomycotina). Microbiology. 140: 2115-2124.

49

Rizzo, D. M., Whiting, E.C., & Elkins, RB. (1998). Spatial Distribution of Armillaria
mellea in Pear Orchards. Plant Disease. 82: 1226-1231.

Schulze, S. & G. Bahnweg (1998). Critical review of Identification Techniques for
Armillaria spp. and Heterobasidion annosum Root and Butt Rot Diseases. Journal
of Phytopathology. 146: 61-72.

Shaw, C. G. 111, & Roth, L.F. (1976). Persistence and distribution of a clone of Armillaria
mellea in a ponderosa pine forest. Phytopathology. 66: 1210-1213.

Shaw, C. G. 111, & Roth, L.F. (1978). Control of Armillaria root rot in managed
coniferous forests. European Journal of Forest Pathology. 8: 163-174.

Sicoli, G., F atehi, J ., & Stenlid, J. (2003). Development of species-specific PCR primers
on rDNA for the identification of European Armillaria species." Forest Pathology.
33: 287-297.

Sierra, A. P., Whitehead, D.S., & Whitehead, MP. (1999). Investigation of a PCR-based
method for the routine identification of British Armillaria species. Mycological
Research. 103: 1631-1636.

Smith A.M., & Griffin, D.M., (1971). Oxygen and the ecology of Armillaria elegans
Heim. Australian Journal of Biological Sciences. 24: 231-262.

Smith, M. L., Bruhn, J .N., & Anderson, J .B. (1992). The fungus Armillaria bulbosa is
among the largest and oldest living organisms. Nature. 356: 428-431.

Smith, M. L., Bruhn, J .N., & Anderson, J .B. (1994). Relatedness and spatial distribution
of Armillaria genets in infecting red pine seedlings. Phytopathology. 84: 822-829.

Smith, M. L., Duchesne, L.C., Bruhn, J .N., & Anderson, J .B. (1990). Mitochondrial
genetics in a natural population of the plant pathogen Armillaria. Genetics. 126:
575-582.

Taylor, J. W, Jacobson, D. J. Kroken, S. Kasuga, T., Geiser, D. M. ,Hibbett, D. S., &
Fisher, M. C. (2000). Phylogenetic Species Recognition and Species Concepts in
Fungi. Fugal Genetics and Biology. 31: 21-32.

Thompson, W. (1984). Distribution, development, and functioning of mycelial cord
decomposer basidiomycetes of the deciduous woodland floor. In: The ecology
and physiology of the fungal mycelium. Eds., D.H. Jennings and A.D.M. Rayner.
Cambridge University Press, Cambridge, United Kingdom. 185-214.

Tommerup, I. C., & Broadbent, D. 1975. Nuclear fusion, meiosis and the origin of
dikaryotic hyphae in Armillariella mellea. Arch. Microbiol. 103: 279-282.

50

Turner, J. A. & Fox, R.T.V. (1988). Prospects for the chemical control of Armillaria
species. In: Brighton Crop Protection Conference: Pests and Diseases. 1. 235-240.

Ullrich, R.C., & Anderson, J .B., (1978). Sex and diploidy in Armillaria mellea.
Experimental Mycology. 2: 1 19-129.

Vance, C. P. & Garraway, MD. (1973). Growth stimulation of Armillaria mellea by

ethanol and other alcohols in relation to phenol concentration. Phytopathology.
63: 743-748.

Volk, T.J. Key to North American Armillaria species. Retrieved May, 16, 2005, from the
Center for Forest Mycological Research, Forest Service, U.S.D.A. at the Dept. of
Botany at the Univeristy of Wisconsin Website:
http://botit.botany.wisc.edu/toms_fungi/armkey.html

Volk, T.J., (2002). The humongous fungus—Ten years later. Inoculum. 53: 4-8.

Volk, T.J., & Burdsall, H.H. Jr. The state of taxonomy of the genus Armillaria. Retrieved
May, 16, 2005, from the Center for Forest Mycological Research, Forest Service,
U.S.D.A. at the Dept. of Botany at the Univeristy of Wisconsin Website:
http://botit.botany.wisc.edu: 16080/toms_fimgi/arm.htrnl

Watling, R., Kile, G.A. & Burdsall, H.H. Jr. (1991). Nomenclature, taxonomy, and
identification. In: Armillaria Root Disease. Washington, DC, United States
Department of Agriculture Forest Service Agriculture Handbook No. 691. 1-9.

White, E. E., Dubetz, C.P., Cruickshank, M.O., & Morrison, DJ. (1998). DNA diagnostic
for Armillaria species in British Columbia: within and between species variation
in the IGS-1 and IGS-2 regions. Mycologia. 90: 125-131.

Worrall, J. J. (1997). "Somatic incompatibility in basidiomycetes." Mycologia. 89: 24-
36.

Worrall, J. J ., Sullivan, K.F., Harrington, T.C., & Steimel, J .P. (2004). Incidence, host
relations and population structure of Armillaria ostoyae in Colorado
campgrounds. Forest Ecology and Management. 192: 191-206.

Wostemeyer, J. & Kreibich, A. (2002). Repetitive DNA elements in fungi (Mycota):
impact on genomic architecture and evolution. Current Genetics. 41: 189-198.

Zomlefer, W.B. (2004) Guild to Flowering Plant Families. The University of North
Carolina Press & Chapel Hill & London, Chapel Hill, NC.

51

CHAPTER II

DISTRIBUTION, IDENTIFICATION, AND POPULATION DIVERSITY OF

ARfiflLLARIA SPP. IN MICHIGAN CHERRY ORCHARDS

52

INTRODUCTION
Armillaria root rot is an important, localized disease affecting Montrnorency tart

cherry and other stone fruit in Michigan. In 1987, Proffer et al. surveyed cherry orchards
in the western cherry—producing region of Michigan for Armillaria infection. In that
study, haploid single basidospore isolates were established, cultured, and identified to
species using sexual mating tests with known tester isolates. Using this method, species
of Armillaria were identified that infected tart cherries in Michigan (Proffer et al. 1987).
I A new Armillaria distribution survey is pertinent because many of the orchards are now
out of production, new orchards have been established, and the methodology used was
limited by the need for sporulating basidiomes to obtain tester isolates. Modern methods,
such as PCR and restriction fragment length polymorphism (RF LP) analysis of the
intergenic spacer 1 (IGS-l) region (Banik et al. 1996; Harrington and Wingfield 1995;
Kim et al. 2000; Sierra et al. 1999; White et al. 1998), have removed the reliance on
basidiomes, and isolates can now be identified to species with mycelium, rhizomorphs,
and basidiomes. This is important because some species of Armillaria causing infection
may have been missed in the 1987 survey if they did not produce mushrooms that year.
The increased flexibility of newer PCR based identification methods and the need to
examine orchard sites currently reporting losses to Armillaria support the need for a
follow up survey.

Many studies in other regions have looked at the distribution of species and the
clonal spread of Armillaria both in orchards and in forests stands (Anderson et. a1. 1979;
Banik et al. 1995, 1996; Bendel 2006; Berthelay and Guillaumin 1985; Blogdett and

Worrall 1992; Guillarnin et al. 1993, 1996; Guillaumin and Berthelay 1990; Harrington

53

and Rizzo 1993; Kable 1974; Kile 1983, 1986; Klein-Gebbinck et al. 1991; Kromroy
2004; Lung-Escarmant and Gayon 2004; Marsh 1952; McLaughlin 2001; Proffer et al.
1987; Prospero et al. 2003; Harrington and Rizzo 1993; Rizzo et a1. 1998; Shaw and Roth
1976; Smith et al. 1992, 1994; Thompson 1984; Ullrich and Anderson 1978; and Worrall
2004). In North America, species distribution studies have mostly been conducted in
forested areas (Blogdett and Worrall 1992; Harrington and Rizzo 1993; McLaughlin
2001). The host range in all three studies found that A. gemina, A. calvescens, A. mellea,
A. gallica, and A. sinapina were almost always found on hardwood species, while A.
ostoyae was the only species predominantly found on conifers. Proffer et al. (1987)
assessed the distribution of Armillaria species in Michigan tart cherry orchards. In this
study, three species of Armillaria were found to be vigorous pathogens to orchard grown
Prunus, including A. ostoyae, A. mellea, and A. calvescens. Although A. ostoyae is a
principal host to conifer, Proffer et al. (1987) found that it is the predominant pathogen to
tart cherry in Leelanau, Grand Traverse, and Benzie counties where cherry production is
greatest and the pathogen is widespread. Many of the clonal studies have focused on the
spread of A. ostoyae in coniferous forests or pine plantations, finding the genets at a
platation site often pre-date the plantation and were established from the existing forest
before conversion (Klein-Gebbinck et al. 1991; Lung-Escarmant and Gayon 2004;
Prospero et al. 2003; Smith et a1. 1994; Worrall 2004). The study by Smith et al. (1994)
examining infected red pine seedlings in Michigan, concluded that based on the estimates
of size and shape, some of the genets were established prior to the time of stand
conversion, and that the largest clone may predate the existing hardwood forest, which

replaced a forest of mostly pine. Rizzo et al. (1998) examined the clonal spread of A.

54

mellea in two pear orchards, previously hardwood forests, in California. In this study,
Rizzo et al. (1998) discovered four genets at one site with three of the genets being over
100 m in length. At the second site five genets were found, however these genets were
smaller, ranging in sizes from 20 to 60 m in length. The conclusions of this study suggest
that based on the size of many of these genets, they were established prior to planting of
the orchards, however the size indicates that expansion has occurred since the pears were
planted. Another study examined the delineation and biology of genets of three A.
ostoyae, A. gemina, and A. calvescens at a mixed conifer-hardwood site and at a spruce-
fir site in New Hampshire (Rizzo and Harrington 1993). The results from this study
found six genets of A. ostoyae at the spruce-fir site, colonizing 34% of the conifers and
13% of the hardwoods. At the mixed conifer-hardwood site A. ostoyae and A. calvescens
were found colonizing both conifers and hardwood hosts, while A. gemina was only
found colonizing only hardwood hosts. Six A. ostoyae genets, two A. calvescens, and
two A. gemina genets were identified using somatic incompatibility tests and isoenzyme
analysis. Revisiting species distribution and examining clonal spread would be the next
step to understanding the Michigan populations of Armillaria currently in the cherry
orchards. Somatic incompatibility tests are the most used method to identify individual
clones of Armillaria, as seen in the many publications (Anderson et al. 1979b; Bendel
2006; Ferguson et al. 2003; Klein-Gebbinck et al. 1991; Lung-Escarmant and Gayon
2004; Prospero et al. 2003; Rizzo et al. 1998; Shaw and Roth 1976, Smith et al.1992;
Smith et al. 1994; Worrall 2004). This method involves pairing test isolates on culture
media and examining the visible interaction of the expanding clones as they make

contact. The formation of a barrage zone along the line of contact between isolates

55

indicates the isolates are different clones. Mating-type allele tests may also be used to
distinguish between individual clones. However, this method may only be used with the
single spore isolates of all mating types. Thus, unless all of the individual clones at the
site produce mushroom, this technique is not useful. Molecular techniques have also
been used to identify clones. Analysis of nuclear DNA using random amplified
polymorphic DNA (RADP) analysis and microsatellites also knovm as simple sequence
repeats (SSRs) correspond well with somatic incompatibility reactions and mating-type
allele tests to show genetic distinction between clones. Langrell et al. (2001) developed a
set of 12 primers for di-nucleotide repeats that could be used to distinguish different
somatic incompatibility groups of Armillaria ostoyae isolates. Worrall et al. (2004) used
two of the (ii-nucleotide primer sets developed by Langrell (2001) and also developed
two primer sets for tri-nucleotide repeats to distinguish clones of A. ostoyae in a Colorado
campground. Although the use of SSRs provides a fast and easy way to identify
individual clones, they still should be backed up with methods such as somatic
incompatibility tests.

The hypotheses of this research were: (1) that new species of Armillaria would be
identified on cherry using molecular identification techniques not reliant on basidiomes
as in the 1987 survey by Proffer et al. (1987); and (2) the diversity of species in an
orchard may be much greater than previously reported. To test these hypotheses a broad
distribution survey of orchards in the cherry producing region of Michigan was done, as
well as a clonal mapping of Armillaria within two Montmorency tart cherry orchards

using somatic incompatibility tests and microsatellite markers.

56

MATERIAL AND METHODS:
SURVEY AND FIELD SAMPLING

Fruit orchards and hardwood deciduous forests in 14 Michigan counties were
examined for the presence of Armillaria spp. in 2005 and 2006 (Table 2-1). In forested
areas, fallen logs and stumps were examined for signs of rhizomorphs and mycelium. In
orchards, the early defoliation of trees, circular pockets of dead and declining trees in the
orchard, and mushrooms characteristic of Armillaria at the base of a tree or stump were
used to identify potential Armillaria sites. Trees or stumps suspected to be infected with
Armillaria were examined by digging approximately 0.5 m around the base. When
present, samples of mycelium and/or rhizomorphs under bark were collected by breaking
off infected tissue with a shovel. The samples were transferred to plastic bags, placed on
ice, and taken back to the laboratory for processing. The type of trees samples were
collected from at the site and the GPS coordinates were recorded when possible.

Samples of unexposed mycelium under the bark were removed by cutting into the
bark with a sterile scalpel to expose mycelium and then small pieces, approximately 2-4
mm in diameter of the freshly exposed mycelium were removed using sterile forceps for
plating on agar medium. Rhizomorphs were cut into 1-2 cm pieces and surface
disinfected in a wash of 10% bleach for 3 min, followed by 30% hydrogen peroxide for 2
min, and a final rinse in sterile deionized water for three min. The samples were blotted
on sterile filter paper before being plated. Samples were plated on 2% malt extract agar
(MEA) (20g/l of malt extract and 20g/l of agar) or 2% water agar. The cultures were kept
in the dark at 25° C. The culture plates were checked daily for signs of contamination
from other microbes. If plates were contaminated, transfers were made from the

Armillaria samples that appeared non-contaminated to a clean 2% MBA plate.

57

Table 2-1: Location, date, and description of sites surveyed for Armillaria root rot

 

 

Site # County GPS Date collected Host

1 Oceana N/A 05-19-05 Tart Cherry

2 Ingham N 42° 43 06-02-05 Hardwood Forest
W 084° 28 06-08-05 (Beech, Oak, Maple,

06-16-05 and Black Cherry)
06-23-05

3 Leelanau N 44° 50 09-09-05 Peach
W 085° 39

4 Van Buren N 42° 19 10-07-05 Plum
W 086° 02

5 Berrien N 42° 08 10-07-05 Tart Cherry
W 086° 15

6 Berrien N 42° 06 10-07-05 Apple
W 086° 17

7 Leelanau N 44° 55 10-14-05 Tart Cherry
W 085° 39

8 Leelanau N 44° 56 10-14-05 Tart Cherry
W 085° 39

9 Leelanau N 44° 56 10-14-05 Tart Cherry
W 085° 37

10 Leelanau N 44° 56 10-14-05 Tart Cheny
W 085° 36

l l Leelanau N 44° 57 10-14-05 Tart Cherry
W 085° 36

12 Leelanau N 45° 03 10-14-05 Tart Cheny
W 085° 34

13 Leelanau N 44° 59 10-14-05 Tart Cheny
W 085° 36

14 Grand Traverse N 44° 46 10-18-05 Tart Cherry
W 085° 44

15 Leelanau N 44° 46 10-18-05 Tart Cheny
W 085° 44

16 Benzie N 44° 39 10-18-05 Tart Cherry
W 086° 04

17 Benzie N 44° 31 10-18-05 Peach
W 086° 07 Tart Cherry

Sweet Chery

 

58

Continuation of Table 2-1: Location, date, and description of sites surveyed for

 

 

Armillaria root rot
Site # County GPS Date collected Host
18 Leelanau N 45° 01 10-18-05 Apple
W 085° 38 Tart Cherry
19 Kent N 43° 09 10-25-05 Apple
W 085° 46
20 Kent N 43° 07 10-25-05 Peach
W 085° 48
21 Kent N 43° 15 10-25-05 Tart Cheny
W 085° 46
22 Clinton N/A 1 1-01-05 Maple, Oak, and Elm
23 lonia N/A 11-01-05 Hardwood Forest
(Beech, Elm, Maple)
24 Grand Traverse N 44° 40 1 1-11-05 Tart Cherry
W 085° 35
25 Antrim N 44° 52 1 1-1 1-05 Tart Cherry
W 085° 25
26 Antrim N 44° 56 1 1-1 1-05 Tart Cherry
W 085° 21
27 Antrim N 44° 56 1 1-1 1-05 Tart Cherry
W 085° 21
28 Antrim N 45° 00 1 1-1 1-05 Tart Cheny
W 085° 21
29 Leelanau N 44° 57 1 1-1 1-05 Tart Cherry
W 085° 36
30 Manistee N 44° 19 06-07-06 Apple
W 086° 14 Sweet Cheny
31 Oceana N/A 06-22-06 Tart Cherry
Plum
32 Oceana N/A 1 1-04-06 Tart Cheny
33 Montcalm N/A 08-04-06 Frasier Fir
34 Oceana N/A 11-04-06 Tart Cheny
35 Oceana N/A 1 1-04-06 Sweet Cherry
36 Tuscola N/ A 10-08-06 Hardwood Forest

 

59

This process was repeated until pure cultures were obtained. Pure cultures were
transferred to 3% MBA and were maintained in an incubator at 25° C in the dark.

During the summer of 2006, intensive sampling was completed in two orchards in
Grand Traverse and Leelanau counties. This was done to examine clonal spread within
Michigan orchards. The two orchards were chosen because they were cleared forested
areas that were planted to Montmorency tart cherry grafted on mahaleb rootstock. The
first orchard (site # 14 from Table 2-1) was approximately 30 years old and was located
along the Leelanau and Grand Traverse county line. The trees in the orchard were
planted 6x6 m apart. The orchard was split into four sections by a single row of poplar
(Populus) trees, with 13 -15 rows per section. Within each section there were circular
pockets where trees had been removed due to infection from Armillaria. There was a
mixed maple (Acer) forest along the south side of the orchard. All 913 trees in the
orchard were examined for the presence of infection from Armillaria. Trees 5 m into the
surrounding forest were also examined for the presence of infection from Armillaria.
Samples were collected and processed as described above.

The second orchard (site # 29 from Table 2-1) surveyed was 30 years old and
located on the Old Mission Peninsula. The trees were planted 3 x 5.4 m apart and the last
14 rows were planted 2.7 x 5.4 m apart. The orchard was divided into four sections,
which was divided by a main gravel road running east to west and the two sections were
divided by a smaller grass path running north to south. The first two sections to the south
of the main gravel road had 16 rows, while the second two sections to the north had 28
rows. There were approximately 50 to 150 trees per row depending on where the row

started and on how many trees had been removed due to Armillaria root rot or other

60

causes such as injury. Forest containing mostly beech (F agus grandifolia Ehrh.) and
maple surrounded the south and west sides of the orchard. The second orchard also had
circular pockets where trees had been removed due to Armillaria infection.
Approximately 50 to 60 trees were lost annually in this orchard to Armillaria infection
according to the grower. All symptomatic trees, trees adjacent to infected trees, and
some asymptomatic trees, totaling approximately 25% of the 4,339 trees in the orchard,
were examined for the presence of Armillaria. Trees 5 m into the surrounding forest
were also examined for the presence of infection by Armillaria. Samples were collected

and processed as described above for the survey.

SPECIES IDENTIFICATION

DNA was extracted from the isolates using one of two methods. For immediate
use, DNA was extracted by scraping a forcep across the edge of a mycelial culture
growing on 3% MBA to collect a 1-2 mm2 piece of mycelium. The tip of the forcep with
the mycelium was then inserted in to a 1.5 ml microcentrifuge tube containing 100 pl 5X
Tris-Borate (TBE) lysis buffer (54g Tris-base, 27.5g boric acid, 20ml 0.5M EDTA [pH
8.0]. The DNA sample was then boiled for 10 min at 100° C. DNA that was going to be
saved for later analysis was isolated by taking 100 mg fresh weight mycelium from a
culture growing on 3% MBA. The mycelium was placed in a 1.5 ml microcentrifuge
tube with two sterilized 3 mm glass beads. The sample was placed in liquid nitrogen for
two minutes, and then macerated using a FastPrep F P120 Homogenizer (Qbiogene Inc.,
Carlsbad, CA 92008) at a speed of 4.5 for 40 s. The last two steps were repeated two-

three times or until samples looked completely macerated. The lysis buffer AP] and the

61

RNase A stock solution from the Qiagen Dneasy® Plant DNA kit (Qiagen Sciences,
Maryland 20874) were added to the macerated sample and homogenized using the
FastPrep FP120 Homogenizer at a speed of 4.5 for 40 3. DNA was then extracted using
the Qiagen Dneasy® Plant DNA kit following the manufactures instructions after the
addition of the AP] buffer and RNase A stock solution. A modification was made at the
final step, where 50 uL of the elution buffer AB (10 mM Tris-Cl , 0.5 mM EDTA; pH
9.0) was added instead of 100 pL to increase the final DNA concentration. The elution
buffer AE was also heated to 65° C before being added to help the long genomic DNA
release from the silica membrane. The two final elutions were combined together in the
same tube. DNA was stored at 4° C.

Established PCR and RFLP protocols for the analysis of the intergenic spacer l
(IGS-1) region (Harrington and Wingfield 1995) were used for species determination.
The primers used to amplify this region were LR12R (Veldman et al. 1981), and O-l
(Duchesne and Anderson 1990), as first recommended by Anderson and Stasovski
(1992). The PCR reaction mixture was modified and included 2.5U of Taq polymerase,
1% 10X PCR buffer, 3.5 mM MgC12, 200 uM dNTPs (Invitrogen, Carlsbad, California
92008), and 0.5 uM of each primer (Integrated DNA Technologies Inc., Coralville, Iowa
52241). DNA was diluted to a 1:10 ratio and 2 pl were added to the reaction mixture for a
final reaction volume of 50 pl. The thermocycler used was a GeneAmp® PCR system
2720(Applied Biosystems, Foster City, CA 94404) and the conditions used were the same
as described by Harrington and Wingfield (1995).

PCR-amplified DNA was first digested with the restriction enzyme Alul, and

subsequently with BsmI and Ndel, as required (New England Biolabs, Ipswich. MA

62

0193 8-2723). The reaction mixture included 2.5 pl of the appropriate buffer supplied
with the enzyme, 2.0 pl of deionized sterile water, and 0.75 pl of the enzyme. Amplified
DNA (12 pl) was added to the reaction mixture for a final reaction volume of 17.25 pl
and allowed to digest for 4-16 h at 37 C° for AluI and NdeI and 65 C° for Bsml.

Amplified DNA and the restriction enzyme fragments of these products were
separated by electrophoreses in 1-2.5% agarose gels in 0.5X TBE buffer to determine the
size of amplification and restriction products. For analysis of the amplified DNA, 2pl of
loading dye (0.25% Bromophenol blue, 0.25% xylene cyanol FF, 15% Ficoll (Type 400;
Pharmacia in water, for a total volume of 50ml) was added to the 12 pl PCR product and
12 pl of this mixture was loaded onto a 1% agarose gel run at 100 V for 1.5 h. For the
restriction products, 2 pl of loading dye was added to the product and 15 pl of this
mixture was loaded to a 2.5% agarose gel run at 60 V for 2.5 h. The banding patterns of
the tested isolates were compared to those of eight different tester isolates of North
American Biological Species (Table 2-2). Fragments longer than 100 bp were scored
both visually and from sequenced isolates using the NEBcutter V2.0 (New England
Biolabs, Ipswich, MA 01938-2723).

Some isolates could not be identified to species using the RFLP protocol of
Harrington and Wingfield (1995). Armillaria clavescens and A. gallica can have the
same AluI banding patterns making them indistinguishable from one another. A. ostoyae
and A. gemina also have the same AluI banding patterns but can usually be distinguished
from each other using different restriction enzymes such as NdeI or Bsml which have
been shown to digest only the PCR product for A. ostoyae isolates. A mutation at these

sites would make the species indistinguishable from one another. Some isolates were

63

identified to species using a modified version of the sexual mating tests described by
Korhonen (1978). The sexual mating test involves pairing of haploid isolates with known
haploid testers on culture media, which are then examined visually for the conversion of
the haploid isolates to diploid form due to sexual compatibility. This test has also been
found to work by pairing diploid isolates converting the haploid testers when sexually
compatible. Using this modified technique, the species of diploid isolates, that could not
be distinguished using the RF LP protocol, was determined. Plugs (5x5 mm) of the
unidentified isolates were placed 10 mm apart from a plug (5x5 mm) of a tester isolate on
a 60 mm by 15mm Petri dish containing 4% MBA amended with 20 g/l glucose and 5 g/l
peptone. Four replications for each tester isolate and unidentified isolate were plated.
Isolates were allowed to grow for four weeks in the dark at room temperature and then

observed for the conversion of the haploid tester to the diploid form.

64

Table 2-2: Tester strains used as controls to compare and identify Michigan isolates

of Armillaria to species

 

 

Name of Strain* N ABS“ Catalog Depositor Location Obtained From
Numbu'

Armillariella mellea 28-4 NABS I 44396 J B Anderson ATCC, Manassas, VA
(AlBl) 20108
Armillariella mellea 28-6 NABS I 44399 JB Anderson ATCC, Manassas, VA
(A1B2) 20108
Armillariella mellea 28-7 NABS I 44401 J B Anderson ATCC, Manassas, VA
(A231) 20108
Armillariella mellea 28-9 NABS I 44403 JB Anderson ATCC, Manassas, VA
(A282) 20108
Armillariella mellea 160-8 NABS 11 52122 AL Jones, TJ Michigan State University

Proffer
Armillariella mellea 160-9 NABS 11 52123 AL Jones, TJ Michigan State University
Proffer
Armillariella mellea 11-1 NABS 111 52580 J B Anderson ATCC, Manassas, VA
20108
Armillariella mellea 11-9 NABS 111 52099 18 Anderson ATCC, Manassas, VA
20108
Armillariella mellea 21-2 NABS 111 52100 JB Anderson ATCC, Manassas, VA
20108
Armillariella mellea 48-1 NABS V 52104 AL Jones, TJ Michigan State University
Ihoflbr
Armillariella mellea 48-6 NABS V 52105 AL Jones, TJ Michigan State University
' Proffer
Armillariella mellea 49-5 NABS V1 52106 J B Anderson ATCC, Manassas, VA
20108
Armillariella mellea 49-8 NABS V1 52582 AL Jones, T] Michigan State University
Proffer '
Armillariella mellea 97-1 NABS V1 52111 AL Jones, TJ Michigan State University
Ihofibr
Armillariella mellea 90-4 NABS V1] 52109 J B Anderson ATCC, Manassas, VA
20108
Armillariella mellea 90-10 NABS VII 521 10 J B Anderson ATCC, Manassas, VA

65

20108

Continuation of Table 2-2: Tester strains used as controls to compare and identify

Michigan isolates of Armillaria to species

 

 

Name of Strain Species Catalog Depositor Location Obtained From
Number
Armillariella mellea 1-137 NABS V11 521 14 JB Anderson ATCC, Manassas, VA
20108

Armillariella mellea 121-2 NABS [X 521 13 AL Jones, TJ Michigan State University
Proffer

Armillariella mellea 139-1 NABS 1X 521 16 AL Jones, TJ Michigan State University
Proffer

Armillariella mellea 140-6 NABS X 521 19 AL Jones, TJ Michigan State University
Proffer

Armillariella mellea 140-9 NABS X 5212] AL Jones, TJ Michigan State University
Proffer

Scytinostroma galaectinum - - TJ Proffer Michigan State University

Smith T1
Polyporus squamosis - - TJ Proffer Michigan State University
Schizophllum commune - - TJ Proffer Michigan State University

 

* Armillariella is a synonym for Armillaria. See the Taxonomy section in Chapter 1 for an explanation.
** NABS = North American Biological Species

66

DNA SEQUENCING OF THE IGS-l REGIONS

A group of isolates were selected, representing all of the distinctive RFLP
patterns, for sequencing. This is because the banding sizes from the restriction digests
did not always add up to the prOper fragment size for the IGS-1 region. This was also
important for the isolates in the A. ostoyae/A. gemina group using the AluI restriction
digest. DNA fragments representing all of the Alul, BsmI, and Ndel banding patterns
were sequenced and compared with one another as well as with sequenced strains on
Genbank. The PCR reaction mixture and the thermocycling conditions were the same as
for species identification with the exceptions of the final reaction volume which was
increased to 100 pl. Loading dye (3 pl) was added to the entire PCR product. The entire
PCR product, 20 pl per well (five wells), was loaded onto a 1% agarose gels which were
run at 100 V for 1.5 h. The DNA was then extracted from the gel using a QIAquick®
Gel Extraction Kit (Qiagen Sciences, Maryland 20874) following the instructions of the
manufacturer. DNA sequencing was performed at the Research Technology Support
Facility (RTSF) at Michigan State University. Sequences using direct PCR and those
obtained from GenBank were analyzed both manually and using Lasergene® version 6

software (DNASTAR Inc., Madison, WI 53705).

SOMATIC INCOMPATIBILITY TESTS
The genet identification of isolates collected from the two orchard sites in the
clonal study was determined using somatic incompatibility tests. Plugs (5x5 mm) of two
different unidentified isolates were plated approximately 10 mm from each other on a 60

mm by 15mm Petri dish containing 4% MBA amended with 20 g/l glucose and 5 g/l

67

peptone. Isolates were then allowed to grow for six to eight weeks in the dark at 25 °C.
After six to eight weeks the cultures were observed for the presence or absence of a
barrage zone to determine the somatic incompatibility (SI) group of each isolate. For the
first orchard, two rounds of pairings were done in order to determine the SI group of each
isolate. Since the second orchard had more isolates, three rounds of pairings were needed
to determine the SI group of each isolate.

Orchard A:

Isolates were paired in all possible combinations. In addition, self-crosses were
included as controls. There were four replications (four plates) for each cross. After this
first round of pairings, the SI group for each isolate could be determined; however, a
second round of parings was used to confirm the results from the first round. In this
second round, a representative isolate from an SI group identified in the first round was
paired with another representative isolate from all other SI groups. Also, each isolate
within an S1 group was crossed with the other isolates within the same SI group to
reconfirm compatibility. Self-crosses were again done as controls and three replications
(three plates) were done for each cross in the second round.

Orchard B:

Because of the greater number of isolates collected from the second orchard, the
isolates were divided into four groups of 20-21 isolates. In the first round of pairings, all
possible crosses within a group were done, as well self-crosses as controls. There were
four replications (four plates) for each cross. In the second round of pairings,
representative isolates from each SI group found in one of the four groups were crossed

with representative isolates from SI group in the three other groups. Self-crosses of each

68

representative isolate were also done as controls. In the second round of pairings, there

were four replications (four plates) for each cross. After the two rounds of pairings were
completed the SI group for each isolate could be determined. However, a third round of
pairings was done to verify the results in the same way that the second round of pairings

was done for the first orchard.

MICROSATTELITE ANALYSIS

Somatic incompatibility groups identified for the A. ostoyae isolates in both
orchards, and A. gemina isolates in orchard B (site # 29) were confirmed to be genetically
distinct using published microsatellite markers, specifically designed for A. ostoyae
(Langrell et al. 2001; and Worrall et al. 2004) (Table 2-3). Two representative isolates
from each SI group in orchard A (site # 14), one representative isolate for each SI group
in orchard B (site # 29), and seven isolates from different orchards were used with the
microsatellite markers to confirm that the A. ostoyae SI groups were genetically distinct
from each other and to show that the two representative isolates in the same SI group had
the same alleles. In addition, the seven other isolates of A. ostoyae from other sites were
tested to see if the microsatellite markers would pick up global diversity between isolates.
The DNA used for the microsatellites was extracted using the second method described
utilizing the Qiagen Dneasy® Plant DNA kit. The DNA was diluted to a 1:10
concentration. The PCR reaction mixture was the same as previously described. The
thermocycling conditions were the same as described by Langrell et al. (2001) with the
annealing temperature ranging from 53-5 8° C depending on the fluorescence labeled

primers (Applied Biosystems, Foster City, CA 94404). The primers were labeled with 6-

69

F AM or HEX at the 5’-end of the forward primer (Table 2-3). The PCR products for
each primer set were purified using a QIAquick® PCR Purification Kit (Qiagen Sciences,
Maryland 20874) following manufacture instructions, and then the products from each set
of primers (5 pl) were combined for an isolate (20 pl final volume). Samples were
submitted to the Research Technology Support Facility (RTSF) at Michigan State
University for analyses via the ABI PRISM® 3100 Genetic Analyzer (Applied
Biosystems, Foster City, CA 94404) and GeneScan® Analysis Software Version 3.7
(Applied Biosystems, Foster City, CA 94404) to identify the sizes of each of the four

loci.

Table 2-3: Microsatellite markers used to genetically confirm SIG’s from the two

intensively sampled orchards

 

Locus Primer Sequences (5’-3’) and Label Observed EM BL Source

Range (bp) No.

 

CAGZS F: 6FAM-CATGACGCCACGGATACCA 356-359 Worrall et
R: TCGCTGACATGTGCCGAGG al. (2004)

CAG77 F: HEX-AGGCTGGCCGAATAGTGAAT 328-349 Worrall et
R: CTGATCTGTGACCTCAAGCA al. (2004)

AOSSR74 F: 6FAM-GCTCACCCTCAAACTTAACA 96-1 14 AJ307595 Langrell et
R: GCAGGGCACAAATGAAACTA al. (2001)

AOSSR84 F: HEX-ACACCACGAGTGCTTCTACTA 128-150 AJ307598 Langrell et
R: GCT TGG TAA TGG GCA GAG al. (200])

 

7O

RESULTS:
SPECIES IDENTIFICATION AND DISTRIBUTION

Amplification of the IGS-1 region and restriction fragments of this region were
obtained for all isolates. The amplified product for the IGS-1 region was 875 bp for all
A. mellea isolates and 920 bp for all other isolates. The amplified product from all
isolates was first digested with Alul, and subsequently with NdeI and BsmI as required.
The size of the resulting fragments was determined using electrophoresis and by
comparison with results based on sequenced isolates using the NEBcutter V2.0 (New
England Biolabs, Ipswich, MA 01938-2723). Using the restriction enzyme Alul, a total
of three patterns were obtained from fragments longer than 100 bp, all of which
corresponded with previously published patterns (Harrington and Wingfield 1995; White
et a1. 1998; Kim et al. 2000) (Fig. 2-1; Appendix A: Table A-l). From 18 isolates the
PCR products yielded fragments of approximately 476 bp and 175 bp, corresponding to
A. mellea RFLP pattern A, as designated by Harrington and Wingfield (1995). From
these 18 isolates, the PCR products from five isolates belonging to the same SI group
from orchard B (site # 29), yielded a strong fragment at approximately 240 bp. This band
wasoften observed as a very faint fragment for the PCR products of other isolates
identified as A. mellea. The fragment of 240 bp was due to partial digestion of the PCR
product and was only observed when running the digested PCR product on agarose gels.
Based on the sequenced data for these isolates, the restriction sites when cut should only
produce fragments of 476 bp and 175 bp. The amount of restriction enzyme used in the
reaction was unable to resolve the partial digestion of the PCR products for these isolates.
There were 104 isolates whose PCR products yielded fragments of approximately 308 bp,

200 bp, and 135 bp. This pattern placed them into either A. ostoyae or A. gemina, as

71

designated by Harrington and Wingfield (1995). From 36 isolates, the PCR products
yielded fiagments at approximately 583 bp and 240 bp, identifying these isolates as either
A. calvescens or A. gallica, as designated by Harrington and Wingfield (1995). Out of
the 36 isolates, the PCR products of 15 isolates also yielded fragments at approximately
400 bp and 200 bp. The fragments of 476 bp and 175 bp were due to partial digestion of
the PCR product observed when running the digested PCR product on agarose gels.
Based on the sequenced data for these isolates the restriction sites only should produce
fragments of 583 bp and 240 bp. The amount of restriction enzyme used in the reaction
was unable to resolve the partial digestion of the PCR products for these isolates.

Isolates placed into the A. ostoyae/A. gemina group by Alul, are usually
distinguished from one another by the restriction enzymes Ndel. The PCR product of
most A. ostoyae isolates yield fragments of approximately 550 bp and 370 bp (Fig. 2-2;
Appendix A: Table A-l). The digestion of the PCR products for A. gemina tester isolates
and previous published reports indicate that no digestion with this enzyme occurred for
this particular species accept in one rare case (Kim et a1 2000). The PCR product of
isolates of A. ostoyae rarely has not digested with this enzyme (Harrington and Wingfield
1995; Kim et a1. 2000). The PCR products of 33 isolates from a total of eight SI groups
from orchard B (site # 29) and two isolates from site # 35 in the A. ostoyae/A. gemina
AluI group did not digest with Ndel. The PCR products of isolates with the AluI banding
patterns for A. mellea pattern A and A.calvescens/A. gallica also did not digest with this
enzyme.

Because the PCR product of some isolates identified as either A. ostoyae or A.

gemina did not digest with Ndel, the restriction enzyme Bsml was used to try and

72

identify the isolates to species. The PCR products of isolates of A. ostoyae have been
reported to digest with this enzyme, whereas A. gemina does not (Harrington and
Wingfield 1995). The digestion of the PCR products with BsmI yielded products at
approximately 620 bp and 300 bp, which distinguished the isolates as A. ostoyae. The
PCR product of 20 of the isolates belonging to four SI groups from orchard B (site # 29)
and the two isolates from site # 35 that did not digest with NdeI also did not digest with
BsmI confirming their identification as A. gemina. The PCR products of two isolates
from one SI group from orchard B (site # 29) that digested with Ndel, did not digest with
BsmI. All PCR product for isolates with the AluI fragment pattern for A. mellea pattern
A digested with BsmI yielding fragments at approximately 550 bp and 325 bp (Fig. 2-3;
Appendix A: Table A-l). Isolates in the A. calvescens/A. gallica group based on AluI did
not digest with BsmI.

While AluI can be used to distinguish isolates in the A. calvescens/A. gallica
group, no restriction enzymes are reported to distinguish between these two species.
Sexual compatibility tests still must be performed to identify these isolates to species.
Sexual compatibility tests were performed for all isolates from the distribution survey in
the A. calvescens/A. gallica group, except for orchard B (site # 29), where representatives
isolate from each of the three SI groups in this group was tested. Based on the crosses all
of these isolates were identified as A. gallica. The tester strain obtained from ATCC,
Armillariella mellea 90-4 (NABS VII) was the source of confirmation and gave the best
results (Fig. 2-4, 2-5; Appendix A: Table A-2).

Sexual compatibility tests were done for a representative isolate from each of the

15 SI groups whose PCR product yielded the AluI banding pattern for A. ostoyae/ A.

73

gemina from orchard B (site # 29), and both isolates from site #35 whose PCR product
did not digest with Ndel and/or Bsml. The sexual mating tests confirmed that each
representative isolate whose PCR product digested with Ndel and/or Bsml from orchard
B (site # 29) to be A. ostoyae. The tester strains obtained from ATCC, Armillariella
mellea 28-6 (NABS I) and Armillariella mellea 28-4 (NABS I) gave the best results.
Some of the A. ostoyae isolates did not convert the tester Armillariella mellea 28-6, most
likely due to incompatible mating types. However, although the tester strain
Armillariella mellea 28-4 (N ABS 1) partially converted to the diploid form with out being
mated the conversion to the diploid form was stronger with the isolates that did not
convert with Armillariella mellea 28-6. The results for the other isolates whose PCR
product did not digest with both Ndel and Bsml were identified as A. gemina. The tester
strain, Armillariella mellea 160-9 (N ABS II) was the source of confirmation and gave the
best results. (Fig. 2-4, 2-5; Appendix A: Table A-3).

Sequencing of the IGS-1 region was also done for representative isolates with
different Alul, Ndel, and Bsml banding patterns to help identify species and determine
fragment sizes from the restriction digests. All sequenced isolates were compared with
sequences on GenBank. Based on the comparison with strains of Armillaria submitted to
GenBank, isolates that were identified as A. mellea were 99% homologous with other A.
mellea isolates on GenBank compared to other species where some A. tabescens isolates
were 95% homologous, and some A. ostoyae and A. gallica isolates were 94%
homologous. Isolates identified as A. ostoyae that digested with both Ndel and Bsml
were 99% homologous with other A. ostoyae isolates and 98% homologous with some A.

gemina isolates on GenBank, whereas isolates that did not digest with Ndel or Bsml were

74

99% homologous with some A. gemina isolates and 98% homologous with some A.
ostoyae isolates. Isolates of A. ostoyae that digested with either Bsml or Ndel were 99%
homologous to isolates of A. ostoyae and A. gemina. Isolates identified as A. gallica
were 99% homologous with other A. gallica isolates and 98% homologous with some A.
clavescens isolates on GenBank.

This new PCR based survey of Armillaria species found in Michigan’s cherry
regions included new areas not surveyed by Proffer et al. (1987). The new orchard areas
surveyed included: the orchard ridge area in Kent County, and Van Buren and Berrien
Counties. Forested areas were surveyed in Clinton, Ionia, Ingham, and Tuscola Counties,
and a sample from a Christmas tree plantation in Montcalm County was added to the
survey. A. ostoyae was the most predominant species found infecting tart cherry and
other stone fruit trees in this survey, especially in the Northwest region. Armillaria
mellea was found infecting cherry trees in Berrien County and occasionally Oceana, and
Leelanau Counties. Armillaria gallica was found on a tart cherry tree in Leelanau
County and on two plum trees in Van Buren County. A. gallica is often noted to act as a
saprophyte in the forest on dead or declining trees, stumps, fallen logs, or in the soil.
However, based on field observations in this survey it appears capable of initiating
disease in orchard grown Prunus spp. Armillaria gemina, never before reported in
Michigan, appeared to be pathogenic on Prunus spp. in Leelanau County on tart cherry

and also on sweet cherry in Oceana County (Table 2-4; Fig. 2-6).

75

Table 2-4: Number of isolates collected from each county, the host, and species

distribution of Armillaria spp. in the cherry producing regions of Michigan

 

 

County Host Number of Species
Isolates
Antrim Tart Cherry 7 A. ostoyae
Benzie Tart Cherry 4 A. ostoyae
Sweet Cherry 1 A. ostoyae
Peach 1 A. ostoyae
Grand Traverse Tart Cherry 22 A. ostoyae
Leelanau Tart Cheny 38 A. ostoyae
20 A. gemina
14 A. mellea
l A. gallica
Peach
1 A. ostoyae
Beech
1 A. ostoyae
2 A. gallica
Maple
l A. gallica
Fallen Log
16 A. gallica
Manistee Sweet Cherry 2 A. ostoyae
Oceana Tart Cheny 4 A. ostoyae
1 A. mellea
Sweet Cheny 2 A. gemina
Plum 1 A. mellea
Montcalm Douglas Fir 1 A. ostoyae
Clinton Fallen Log 2 A. gallica
Ingham Fallen Log 6 A. gallica
Ionia Fallen Log 5 A. gallica
Kent Apple 0
Peach 0
Tart Cherry 0

 

 

 

 

76

Continuation of Table 2-4: Number of isolates collected from each county, the host, and

species distribution of Armillaria spp. in the cherry producing regions of Michigan

 

 

County Host Number of Species
Isolates
Berrien Tart Cherry 2 A. mellea
Apple 0
Van Buren Plum 1 A. gallica
Tuscola Fallen Log 2 A. gallica

 

 

 

 

See Appendix I for complete data for all isolates including; site, tree type, banding size for the IGS-1
region, RF LP fragment sizes for Alul, Bsml, and Ndel, mating compatibility tests, and species.

77

Figure 2-1.
AluI banding patterns observed for Michigan Isolates of Armillaria spp.

1 2 3 4 5 6

’
v
—
’
’
—
-
-
-

l r minim l

.,"
I
Ill
‘0

 

Fig. 2-1: RFLP analysis of the IGS-1 region amplified from Armillaria species. The DNA products
were digested with Alul. Lanes 1 and 6, 100 bp ladder; lanes 2, A. ostoyae /A. gemina; lanes 3 A.
calvescens /A. gallica; lanes 4-5 A. mellea.

78

Figure 2-2.

Ndel banding patterns observed for Michigan Isolates of Armillaria spp.

1 2 3 4 5 6

 

Fig. 2-2: RFLP analysis of the IGS-1 region amplified from Armillaria species. The DNA products were
digested with Ndel. Lanes 1 and 6, [KB plus DNA ladder; lanes 2 and 3, A. ostoyae; and lanes 4 and 5, A.
gemina.

79

Figure 2-3.

Bsml banding patterns observed for Michigan Isolates of Armillaria spp.

 

Fig. 2-3: RFLP analysis of the IGS-1 region amplified from Armillaria species. The DNA products were
digested with Bsml. Lanes 1 and 6, lKB plus DNA ladder; lane 2, A. gemina; lane 3, A. mellea; and lanes 4
and 5 A. ostoyae.

80

Figure 2-4
Sexual compatibility test results for species identification of Armillaria isolates,

using haploid tester isolates

e. V i s ' ,
. . ' hfl‘» _
43113;; "leeks 23' .. “

 

Fig. 2-4: Sexual compatibility tests: Starting from the top left: 90-4 x 90-4 (self cross); 90-4 x 137-1
(compatible); 137-1 x 137-1 (self cross); 28-6 x 28-6 (self cross); 28-6 x 160-9 (incompatible); 160-9 x
160-9 (self cross)

81

Figure 2-5

Sexual compatibility test results for species identification of Armillaria isolates,

using haploid testers and unknown diploid isolates

 

Fig. 2-5: Sexual compatibility tests: Starting from the top left: RR R4-T57 x RR R4T57 (self cross); RR
R4T57 x 90-4 (compatible); 90-4 x 90-4 (self cross); LR R14T5 x LR R14T5 (self cross); LR R14T5 x
160-9 (incompatible); 160-9 x 160-9 (self cross)

82

Figure 2-6.

Distribution map of species of Armillaria in Michigan’s cherry producing regions

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

0| 0 "r
OCEANA A
”A...
“'5'" A A l __
'°"" m“ r
l l I»; l
I
A
J3" _ Legend
* | Aostoyae
. I l ‘ O Agemlna
‘ 7 . Amellea

 

 

 

 

A A. gallica

 

83

 

SOMATIC INCOMPATIBILITY AND MICROSATELLITE RESULTS:

The observed range of allele sizes for the four loci (CAG 25, CAG 77, AOSSR74,
and AOSSR84) corresponded with previous reports by Langrell et al. (2001) and Worrall
et a1 (2004). Each compatibility group of the A. ostoyae and A. gemina strains had a
unique set of alleles for the four loci. However, similar alleles for each of the four loci
were found at all the sites and the alleles of A. gemina were also the same sizes as many
of the A. ostoyae strains. The similarity in allele size could be explained by the close
relationship between the two species compared to the other species of Armillaria. PCR
product for other species were preliminarily tested with each primer set and found that
the primers for the loci CAG77 and AOSSR84 did not anneal to the other species,
whereas strains of A. ostoyae and A. gemina always had PCR products for all four primer
sets.

In orchard A (site # 14) a total of 22 isolates was collected and identified as A.
ostoyae. From these 22 isolates, six genetically distinct genets were found using both
somatic incompatibility tests and microsatellite analysis (Table 2-5; Fig. 2-7, 2-8). The
alleles for orchard A (site # 14) were approximately 348, 353, 355, and 356 for locus
CAG25; 322, 324, 327, 330, and 336 for locus CAG77; 126, 128, 130, 132, 135, and 137
for locus AOSSR84; and 82, 90, 92, 94, 96, 98, and 100 for locus AOSSR74. The
isolates were homozygous or heterozygous for the loci CAGZS and CAG77, and they
were all heterozygous for the locus AOSSR84 and AOSSR74.

Orchard B (site # 29) a total of 81 isolates were collected. From these 81 isolates
26 isolates representing 11 genets were identified as A. ostoyae, 20 isolates representing

four genets were identified as A. gemina,] 5 isolates representing four genets were

84

identified as A. mellea, and 20 isolates representing three genets were identified as A.
gallica (Table 2-5; Fig. 2-7, 2-9). Microsatellites were used to confirm genetic diversity
between clones of A. ostoyae and A. gemina in orchard B (site # 29). The alleles for A.
ostoyae isolates were approximately 348, 351, 353, and 356 for locus CAG25; 322, 327,
330 and 336 for locus CAG77; 122, 126, 128, 130, 132, 135, and 137 for locus
AOSSR84; and 82, 86, 88, 90, 92, 94, 96, 98, 100, 102, 104, and 106 for locus
AOSSR74. The isolates were homozygous or heterozygous for the four loci. The alleles
for the A. gemina isolates were approximately 348 and 351 for locus CAGZS; 330 and
346 for locus CAG77; 126, 128, 130, and 132 for locus AOSSR84; and 90, 92, 94, 96,
100, and 104 for locus AOSSR74. The isolates were homozygous or heterozygous for
the loci CAG25 and CAG77, and they were all heterozygous for the locus AOSSR84 and
AOSSR74.

Seven isolates from other orchards were analyzed to see if the markers could
distinguish genetic diversity at a more global level. The alleles observed at these seven
different orchards were approximately 342, 344, 346, 348, 353, and 356 for locus
CAGZS; 322, 327, 330, and 336 for locus CAG77; 120, 122, 124, 126, 128, 130, 132, and
134 for locus AOSSR84; and 80, 82, 84, 86, 88, 90, 92, 94, 96, 98, 100, and 102 for locus
AOSSR84 (Table 2-5). The isolates were homozygous or heterozygous for the loci
CAG77, and AOSSR74, and they were all heterozygous for the loci CAG25 and
AOSSR84. The isolates were homozygous or heterozygous for the loci CAG25 and

CAG77, and they were all heterozygous for the locus AOSSR84 and AOSSR74.

85

Table 2-5: Somatic incompatibility groups and microsatellite results for orchard A

(site # l4), orchard B (site # 29), and seven miscellaneous strains of A. ostoyae

collected from other sites

 

 

Site SI group # of CAGZS CAG77 AOSRR84 AOSSR74
Isolates
in 81 group

14 Ostl-1 3 356 327 126/137 92/94/96

14 Ost2-1 3 353 322/324 128/137 98/100

14 Ost3-1 3 353/356 322/324 135/137 96/100

14 Ost4-1 4 348/356 327/336 126/130/135/137 90/96

14 Ost5-l 4 353 327 128/130/132/135 82/92/94/96/98
14 Ost6-1 5 348/3 56 3 30 126/130/137 92/94/96/98

29 Ostl-2 1 351/353 327/330 122/137 82/86/92/104/106
29 Ost2-2 4 356 336 122/126/128/130 86/90/94

29 Ost3-2 1 356 327/336 122/128 94/96/98/100
29 Ost4-2 4 351 3228 32 128/132 90/92/94/98/104
29 Ost5-2 2 348/356 327 126/128 86/92/94/98
29 Ost6-2 5 353/356 330/336 132/137 82/88/92

29 Ost7-2 2 351 332 128/130 88/90/92/94/98
29 Ost8-2 2 353/356 332 132/135/137 92/100/102/104/106 ‘
29 Ost9-2 3 351/353 330 132 92

29 Ost10-2 5 351/353 327 126/128/135/137 82/92/98/102
29 Ost11-2 2 351/356 327/336 128/137 90/94/100/102/104
29 Gem 1-2 14 348/351 330 126/128 90/92/94/96
29 Gem2-2 1 348 330/346 128/130/132 90/92/94/96/100
29 Gem3-2 l 351 330/346 128/130/132 90/92/94/96/104
29 Gem4-2 1 348 330/346 128/130/132 90/92/94

29 Me11-2 8 - - - -

29 Me12-2 5 - - - -

 

Continuation of Table 2-5: Somatic incompatibility groups and microsatellite results for

orchard A (site # 14), orchard B (site # 29), and seven miscellaneous strains of A. ostoyae

collected from other sites

 

 

Site SI group # of CAGZS CAG77 AOSRR84 AOSSR74
Isolates
in 81 group

29 Mel3-2 1 - - - -

29 Mel4-2 1 - - - -

29 Cal 1 -2 1 - - - -

29 Ca12-2 8 - - - -

29 Cal3-2 1 1 - - - -

3 .PT1T2 - 342/356 330/336 124/ l 30/‘1 32/1 34 94/98/ 100

17 CT12T1 - 344/356 330 126/ 128/ 130 90/94/100/102
26 CT 1 9T2 - 356 322/327 124/128 80/84/86/90/92/102
30 SC2T1 - 356 322/327 130/ 1 32 82/92/94/96/98
32 CT23T1 - 346/ 356 330 126/132 84/88/90/102
33 XTlTl - 353/356 327/330 132/134 84/92/98/ 100
35 SC3T1 - 344/348 327/330 120/122/124/126/128 80/90/92/94/100

 

See Appendix 2 for data for all isolates.

87

Figure 2-7.
Somatic incompatibility tests used to define genets of Armillaria

in orchard A (site #14) and orchard B (site #29)

Compatible reaction

   

Incompatible reaction

   

Fig. 2-7: Somatic Incompatibility Tests: Plugs (5x5 mm) of an unidentified isolate plated
approximately 10 mm from each other on a 60 mm by 15mm Petri dish. Starting from
the top left: plates 1, 3, 4, and 6, self-crosses or controls; plate 2, compatible reaction; and
plate 5, incompatible reaction, seen by the presence of a barrage zone.

88

Figure 2-8.
Distribution of A. ostoyae genets in orchard A (site # 14) based on results from

somatic incompatibility tests and microsatellite markers

 

fl

 

 

0 15 30 45 60 Meters
Ll—L_L_I_I

 

 

 

Fig. 2-8: First orchard (site # 14) distribution of six genets of Armillaria ostoyae: The red
circles represent trees that isolates were collected from and the lines connecting the
circles represent the area of possible spread of an individual genet. The green lines
represent four single rows of poplar trees that separated the orchard into four sections and
the surrounding stand of maple trees.

89

Figure 2-9.
Distribution results of Armillaria genets for orchard B (site #29) based on results

from somatic incompatibility tests and microsatellite markers

 

 

0 15 30 45 60 Meters
|_r_I_I_I._l

 

 

 

Fig. 2-9: Second orchard (site # 29) distribution of genets of Armillaria spp.: The circles
represent trees that isolates were collected from and the lines represent the area of
possible spread of an individual genet. The red represents A. ostoyae; the brown
represents A. gemina; the blue represents A. mellea; and the green represent A. gallica.
The black represent the edge of the surrounding forest.

90

DISCUSSION:

The amplification and RF LP analysis of the IGS-1 region using Alul, Bsml, and
NdeI digests to identify isolates of Armillaria in this survey yielded RFLP patterns
consistent with previous published reports (Harrington and Wingfield 1995; Kim et al.
2000; White et al. 1998). This method of identification was not reliant on basidiomes,
like the 1987 survey (Proffer et al. 1987) and also allowed for faster identification than
the sexual compatibility test. However this molecular method could not differentiate A.
calvescens from A. gallica. Therefore, there is still a reliance on the sexual compatibility
tests to confirm the species of Armillaria with the AluI banding patterns of A.
calvescens/A. gallica.

Some isolates recovered in the study had RF LP patterns for A. gemina, which has
never been reported in Michigan. Representative isolates placed in the A. ostoyae/A.
gemina group with AluI were paired in sexual compatibility tests to confirm the species
identification. The isolates with the RFLP patterns for A. gemina, whose PCR product
did not digest with Ndel and Bsml, were confirmed to species using the sexual
compatibility tests. The sexually compatibility tests were consistent with the observed
RFLP patterns for the PCR products of A. ostoyae and A. gemina confirming that RFLP
analysis is enough to identify between these two species.

Although sexual compatibility tests were able to distinguish the species of tested
isolates, most of the established tester strains used for the sexual compatibility tests did
not prove to be very efficient. Most of the tester strains were obtained from ATCC, and
many were already in the diploid form or converted to the diploid form without being

mated. Also some tester strains that maintained the haploid phenotype did not convert

91

even with the tester strains of the same species (Appendix A: Table A-2; Table A-3). In
the future, the reliance on a few tester strains for each species using the ATCC as a
source will make it difficult and perhaps impossible to identify some isolates of
Armillaria to species. Armillaria has a bifactorial mating system and if the isolates being
tested are incompatible with the mating type of the only reliable tester the species can not
be determined. If this method is to be used in the future to identify unknown isolates to
species, new testers for all of the biological species should be collected. These testers
were first used by Anderson and Ullrich (1979), so the tester strains may have mutated,
degenerated, or lost viability since they are almost 30 years old.

Further support for species identification can be seen from the sequencing results
of select isolates, which corresponded with both the RFLP analyses and the sexual
compatibility tests. Sequenced isolates were compared to other isolates submitted to
Genbank and were found to be most closely related to isolates of the same species.
However, isolates identified as A. ostoyae that digested with only one of the two
enzymes, Bsml or Ndel, were 99% homologous to isolates of A. ostoyae and A. gemina.
More isolates, should be tested with this method of identification, since many of the
species are closely related, and should still be supported by other identification methods
such as RFLP analysis and the sexual compatibility tests.

The results from this distribution survey correspond in most regards with the 1987
survey (Proffer et al. 1987). A. ostoyae was again found to be the predominant species
infecting tart cherry in the Northwest cherry producing region. Armillaria mellea was
still found to be more predominant as a pathogen on tart cherries in the southern regions

of Michigan. In a new finding of the current survey, A. mellea was found further north

92

than previously reported. In the 1987 survey (Proffer et al. 1987), the farthest north that
A. mellea was reported was in Manistee County on one oak tree. However, isolates of A.
mellea were found farther north on tart cherry in orchard B (site # 29) in Leelanau
County on the Old Mission Peninsula.

Armillaria gallica was found at orchard B (site #29) in Leelanau County on a tart
cherry tree and in the forest as a saprophyte. This species was also found on a plum tree
at site # 4 in Van Buren County. For the most part, A. gallica is considered a saprophyte
and not pathogenic, however, based on field observation in both orchards it appeared to
be acting as an opportunistic pathogen to orchard grown Prunus spp. To confirm the
pathogenicity of A. gallica to Prunus spp. greenhouse inoculation trials should be done.

Isolates of A. gemina were found at orchard B (site #29) on tart cherry.
Armillaria gemina was also identified from site # 35 in Oceana County on sweet cherry.
Armillaria gemina is usually not known as a virulent pathogen. However, based on field
observation at these two sites, A. gemina appeared to be the primary agent of disease and
decline to many of the orchard trees. The pathogenicity of A. gemina to orchard grown
Prunus spp. would not be surprising, since unlike A. ostoyae which tends to be found on
conifers, A. gemina has only been reported on hardwood species (Blogdett and Worrall
1992; Harrington and Rizzo 1993; McLaughlin 2001; Rizzo and Harrington 1993; Volk
and Burdsall). Based on the field observations and host specificity, pathogenicity tests
should be done to confirm that A. gemina can be pathogenic to orchard grown Prunus
spp.

The isolates of A. gemina, A. mellea, and A. gallica found at orchard B (site # 29)

were not found when conducting the broad survey, where only a few isolates were

93

collected from each site. However, when this site was extensively surveyed isolates from
all four species, as including A. ostoyae, were collected. The extensive survey from
orchard B (site # 29) may indicate that other orchards are also diverse in the species of
Armillaria infecting trees, however due to chance and because A. ostoyae is the most
dominant species infecting cherry other species were not collected when broadly
surveying an orchard site. This finding can also support the idea that some species may
have not been identified in the 1987 survey by Proffer et al., which relied on the
production of basidiomes. Species not producing basidiomes in an orchard at the time of
the survey would have been missed.

Based on somatic incompatibility results and microsatellite results, two
intensively surveyed orchards were occupied by multiple Armillaria genets.
Identification of isolates using established microsatellite markers for A. ostoyae and A.
gemina isolates prove to be a reliable and faster method to distinguish individual genets
within Michigan populations. It appears that this method of identification can remove the
reliance on somatic incompatibility identification tests for these Michigan populations.
However, when initially looking at new populations, other available markers should be
evaluated for the specific population to identify which markers show the best variation
for the specific loci. Also, when investigating a new population other identification
methods should still support the use of microsatellite markers.

The genets from both orchards occupied rather small and discrete areas. In
orchard A (site # 14) there seemed to be no division or fragmentation between the genets.
However, this may be due to the low number of sample found at the site. The low

number of samples is due in part to the age and the lack of tree maintenance at the site,

94

resulting in fewer trees to sample. Many of the trees within infection pockets were no
longer present and the existing stumps that looked as though they had been invaded by
Armillaria, due to the presence of dried mycelial fans, wood decay, and/or the presence
of zone lines, were too old to collect viable isolates. Many of the stumps were
decorticated or secondary fungi had moved into the stumps. However, it is difficult to
determine if the existing infestation occurring at this cherry site was subsequent to
orchard establishment. Based on the radial grth rate of A. ostoyae of approximately 1
m per year for young conifer stands (Peet et al. 1996; and Shaw and Roth 1976) it
appears that infestation occurred prior to orchards establishment. This rate of grth is
supported by largest genet at the site, which can be estimated to be around 40 to 100
years in age. Many of the smaller genets may have been established after the orchards
establishment. However based on sample size it is hard to determine if fungal
establishment occurred subsequent to orchard establishment. Based on field observation
and the pocket sizes of areas missing trees, most of the infetation probably occurred prior
to orchard establishment.

At orchard B (site # 29) many of the genets appear to occupy the same space;
suggesting that multiple ramets are at the site. A rarnet is a term to describe a genet that
has been fragmented into two or more pieces; usually due to invasion by other genets.
Ramats could explain why many of the genets found at this site appear to overlie one
another when mapped out. The mosaic pattern of genets at this orchard correspond with
previous studies that looked at clonal spread in forests (Rizzo and Harrington 1993;
Rizzo et al. 1995; Worrall 1994; and Worrall et a1. 2004). This correspondence with

forest populations indicates the probability that these genets are left over from the

95

existing forest and simply moved to the cherry when the orchard was planted. This can
also be supported by the radial spread of the largest genet which at the rate of 1 m per
year (Peet et al. 1996; Shaw and Roth 1976) can be estimated at approximately 70 to 140
years in age. However, the multiple genets infecting a single tree could suggest that these
genets occurred after orchard establishment due to new infection from basidiospores.

The population dynamics at orchard B (site # 29) could be a combination of old genets
established prior to orchard establishment as well as new infection seen by a genet

infecting a single tree.

96

REFERENCES

Anderson, J.B., Ullrich, R.C., Roth, L.F., & Filip, G.M., (1979). Genetic identification of
clones of Armillaria mellea in coniferous forests in Washington. Phytopathology.
69:1109-1111.

Anderson, J .B., & Stasovski, E. (1992). Molecular phylogeny of Northern hemisphere
species of Armillaria. Mycologia. 84: 505-516.

Banik, M.T., Paul, J .A., & Burdsall, H.H. Jr. (1995). Identification of Armillaria species
from Wisconsin and adjacent areas. Mycologia. 87: 707-712.

Banik, M.T., Volk T.J., & Burdsall, H.H. Jr. (1996). Armillaria species of the Olympic
Peninsula of Washington state, including confirmation of North American
biological species XI. Mycologia. 88: 492-496

Bendel, M., Kienast, F. & Rigling, D. (2006). Genetic population structure of three
Armillaria species at the landscape scale: a case study from Swiss Pinus mugo
forests. Mycologial Research. 110: 705-712.

Berthelay, S., & Guillaumin, J .J . (1985). Geographic-distribution of the incompatibility
factors of the diploid Basidiomycete Arrnillaria-obscura (Secretan) Herink.
Cryptogamie Mycologie. 6: 185-196.

Blodgett, J. T. & Worrall, J .J . (1992). Distributions and Hosts of Armillaria species in
New York. Plant Disease. 76: 166-170.

Duchesne, L.C., & Anderson, J .B. (1990). Location and direction of transcription of the
SS r RNA gene of Armillaria. Mycological Research. 94: 266-269.

Guillaumin, J.-J., Anderson, J .B., Legrand, P., Ghahari, S., & Berthelay, S. (1996). A
comparison of different methods for the identification of genets of Armillaria spp.
New Phytologist. 133: 333-343.

Guillaumin, J .J ., & Berthelay, S. (1990). Comparison of 2 methods for the identification
of the clones of the parasitic basidiomycete Armillaria obscura (=A-0st0yae).
European Journal of Forest Pathology. 20: 257-268.

Guillaumin, J .-J ., C. Mohammed, C, Anselmi, N., Courtecuisse, R., Gregory, S.C.,
Holdenridder, 0., Intini, M., Lung, B., Marxmuller, H., Morrison, D., Rishbeth,
J ., Termorshuizen, A.J., Tirro, A, & Van Dam, B. (1993). Geographical
distribution and ecology of the Armillaria species in western Europe. European
Journal of Forest Pathology. 23: 321-341.

Harrington, T. C. & Rizzo, D.M. (1993). Identification of Armillaria species from New
Hamshire. Mycologia. 85: 365-368.

97

Harrington, T. C. & Wingfield, 3D. (1995). A PCR-based identifcation method for
species of Armillaria. Mycologia. 87: 280-288.

Kable, P. F. (1974). Spread of Armillariella sp. in a peach orchard. Transactions of the
British Mycological Society. 62: 89-98.

Kile, GA. (1983). Identification of genotypes and clonal development of Armillaria
luteobubalina Watling & Kile in eucalypt forest. Australiasian Journal of Botany.
31: 657-671.

Kile, GA. (1986). Genotypes of Armillaria hinnulea in wet sclerophyll eucalyptis forest
in Tasmania. Transactions of the British Mycological Society. 87: 312-314.

Kim, M.-S., Klopfenstein, N.B., McDonald, G.I., Arumuganathan, K., & Vidaver, AK.
(2000). Characterization of North American Armillaria species by nuclear DNA
content and RFLP analysis. Mycologia. 92: 874-883.

Klein-Gebbinck, H. W., Blenis, P.V., & Hiratsuka, Y. (1991). Clones of Armillaria
ostoyae and the pattern of infected juvenile lodgepole pine in Alberta, Canada.
European Journal of Forest Pathology. 21: 260-266.

Korhonen, K. (1978). Infertility and clonal size in the Armillariella complex. Karstenia.
1 8: 3 1-42.

Kromroy, K. W. (2004). Identification of Armillaria species in the Chequamegon-Nicolet
National Forest. Res. Note NC-3 88. St. Paul, MN: US. Department of
Agriculture, Forest Service, North Central Research Station. 10 p.

Langrell, S. R. H., Lung-Escarmant, B., & Decroocq, S. (2001). Isolation and
characterization of polymorphic simple sequence repeat loci in Armillaria
ostoyae. Molecular Ecology Notes. 1: 305-307.

Lung-Escarmant, B. & D. Guyon, D. (2004). Temporal and Spatial Dynamics of Primary
and Secondary Infection by Armillaria ostoyae in a Pinus pinaster Plantation.
Phytopathology. 94: 125-131.

Marsh, R. W. (1952). Field observations on the spread of Armillaria mellea in apple
orchards and in a blackcurrant plantation. Trans. Br. mycol. Soc.: 201-207.

McLaughlin, J. A. (2001). Distribution, hosts, and site relationships of Armillaria spp. in
central and southern Ontario. Canadian Journal of Forest Research. 31: 1481-
1490.

Peet, F .G., Morrison, D.J., & Pellow, K.W. (1996). Rate of spread of Armillaria ostoyae

in two Douglas-fir plantations in the southern interior of British Columbia.
Canadian Journal of Forest Research. 26: 148-151.

98

Proffer, T. J ., Jones, A.L., & Ehret, GR. (1987). Biological species of Armillaria isolated
from sour cherry orchards in Michigan. Phytopathology. 77: 941-943.

Prospero, S., Rigling, D., & Holdenridder, O. (2003). Population structure of Armillaria
species in managed Norway spruce stands in the Alps. New Phytologist. 158:
365-3 73.

Rizzo, D. M. & Harrington, T.C. (1993). Delineation and biology of clones of Armillaria
ostoyae, A. gemina, and A. calvescens. Mycologia. 85: 164-174.

Rizzo, D. M., Whiting, E.C., & Elkins, RB. (1998). Spatial Distribution ofArmiIlaria
mellea in Pear Orchards. Plant Disease. 82: 1226-1231.

Shaw, C. G. 111, & Roth, L.F. (1976). Persistence and distribution of a clone of Armillaria
mellea in a ponderosa pine forest. Phytopathology. 66: 1210-1213.

Sierra, A. P., Whitehead, D.S., & Whitehead, M.P. (1999). Investigation of a PCR-based
method for the routine identification of British Armillaria species. Mycological
Research. 103: 1631-1636.

Smith, M. L., Bruhn, J .N., & Anderson, J .B. (1992). The fungus Armillaria bulbosa is
among the largest and oldest living organisms. Nature. 356: 428-431.

Smith, M. L., Bruhn, J .N., & Anderson, J .B. (1994). Relatedness and spatial distribution
of Armillaria genets in infecting red pine seedlings. Phytopathology. 84: 822-829.

Smith, M. L., Duchesne, L.C., Bruhn, J .N., & Anderson, J .B. (1990). Mitochondrial
genetics in a natural population of the plant pathogen Armillaria. Genetics. 126:
575-582.

Thompson, W. (1984). Distribution, development, and functioning of mycelial cord
decomposer basidiomycetes of the deciduous woodland floor. In: The ecology
and physiology of the fungal mycelium. Eds., D.H. Jennings and A.D.M. Rayner.
Cambridge University Press, Cambridge, United Kingdom. 185-214.

Ullrich, R.C., & Anderson, J.B., (1978). Sex and diploidy in Armillaria mellea.
Experimental Mycology. 2: 1 19-129.

Veldman, G.M., Klootwijk, J., de Regt V.C.H.F., & Rudi, RI. (1981). The primary and
secondary structure of yeast 26S rRNA. Nucleic Acids Research. 9: 6935-6952.

White, E. E., Dubetz, C.P., Cruickshank, M.O., & Morrison, DJ. (1998). DNA diagnostic

for Armillaria species in British Columbia: within and between species variation
in the IGS-1 and IGS-2 regions. Mycologia 90: 125-131.

99

Worrall, J. J ., Sullivan, K.F., Harrington, T.C., & Steimel, J .P. (2004). Incidence, host
relations and population structure of Armillaria ostoyae in Colorado
campgrounds. Forest Ecology and Management. 192: 191-206.

100

FUTURE WORK

This research has expanded the fundamental understanding of the population
diversity of Armillaria spp. in Michigan, first initiated by Proffer et al. (1987). More
orchards should be intensively surveyed to see if other orchards have the same diversity
in species seen in orchard B (site # 29), or if a single species usually dominates an
orchard site. Also a particular orchard could be studied over many years, particularly
orchard B (site # 29), to examine the radial rate of spread in cherry, and to examine the
dynamics between the different species of Armillaria found at the site. If this orchard.
was studied over more years, it could be possible to see if new infection is occurring at
the site, as thought possible, based on genets infecting a single tree.

From the large collection of isolates from the survey, pathogenicity screening
could be done to rate the virulence of individual isolates. Isolates of A. gemina and A.
gallica, found on cherry, should be tested for pathogencity to cherry in order to prove
Koch’s postulate and that these species are indeed pathogenic to cherry. Since orchard
site in Michigan are being converted to vineyards pathogenicity screening of the other
species found on Prunus in Michigan, besides A. ostoyae and A. mellea, should be done
including A. gemina, A. calvescens (reported by Proffer et al. 1987), and A. gallica. This
also would be important since one of the species found on cherry, A. gallica, has been
reported on grape by Aguin-Casal et a]. (2004).

Another interesting study would be to look more closely at the life cycle of A.
ostoyae. It has been shown that both A. gallica and A. tabsecens go through two
diploidizations and two haploidization cycles prior to formation of the basidia and are

also genetic mosaics. It would be interesting to see if this is also true for other species of

101

Armillaria, specifically A. ostoyae. Many of the strains collected from the survey of A.
ostoyae converted back and forth between the diploid and haploid phase when cultured
from the same source. From these isolates, basidiome development could be induced to

study the nuclear state of single cells. Also the nuclear state of cells from basidiomes

from nature could be examined.

102

 

 

 

 

 

 

 

assess a. SN 5% EN 52 2. .SN .2: oNo 58a NEE N
. SN

85% .w - - .SN .84 .Nwm o8 2%: _-mo-NNe-3m N
SN

85% e. - - .oeN .84 .me ONO 2%: Nasseém N
SN

8.5% .w - - .oeN .84 .Nwm ONo NS :23 Isaiam N
SN

85% .w - - .SN .84 .NR ON... won 5:5 Néseém N
SN

8.5% .w - - .oeN .84 .23 80 €25 seam 4-8-3-23 N
EN

833... .w - - .SN .84 .Sm ONO E26 seem 78.35%. N

ssass .w 8m 58 EN 52 2. SN .8... CNN. Page sow 78-2-9800 _

as
.833— a
3:25 as .58 as .82 as .2... Two. 2.: 8F 328. am

 

 

 

 

 

 

mas—es 3.255»an cogs—:33 he 5235833.. 83on ES .338.— «new...

«Emu—5 553.58.. 58»?— TmU— 2: new on? mum—.53 Jab 3.: .5539— "—-< 03:.

mHADmm—y— ZO—H<UE—PZm—G— mm—Um—mm

"4.. X—GZHn—m<

103

 

 

 

 

 

 

 

 

 

 

 

 

s98.8 .w OOO .ONO ONO .OOO O2 .OON .OON ONO E26 sow _O_-_ S
asses .w OOO .ONO ONO .OOO O2 .OON .OOO ONO 926 saw OO; 1
sexes .w OOO .ONO ONO .OOO O2 .OON .OOO ONO N56 58 NNOB 2
scene .w OOO .ONO ONO .OOO O2 .OON .OOO ONO 926 58 EOE 2
means s. OOO .ONO ONO .OOO O2 .OON .OOO ONO E26 58 N55 N_
Osees .w OOO .ONO ONO .OOO OO. .OON .OON ONO Ego sow EEO N_

- - - - . EEO Sam 25 Z _ _
eases .w OOO .ONO ONO .OOO O2 .OON .OON ONO E26 sow COD 2
msees .w OOO .ONO ONO .OOO O2 .OON .OOO ONO 920 Bow ONOB O
miss .w OOO .ONO ONO .OOO ON. .OON .OOO ONO E26 58 EOE O
3.8% .w OON .ONO ONO .OOO O: .OON .OOO ONO €26 sow COB O
38.8 .w OOO .ONO ONO .OOO O2 .OON .OON ONO N55 5% COB O
asses .w OOO .ONO ONO .OOO O2 .OON .OOO ONO E26 BOO ENE N
space .V OOO .ONO ONO .OOO ON. .OON .OOO ONO E25 sow ENE N

- - - - - o_an< 0:02 0
see: .w - - ON_ .ONO ONO E26 sow Ne _ so O
uses .4. - - ON. ONO ONO E26 sow E5 O
8st .w - - OON .OOO ONO as: E: 4

 

 

 

 

 

 

 

 

 

 

 

104

 

 

313.3 .
enacts .
Sukarno .
messed .
453.28 .
sundae .
enacts.
EASE .
Essie .
makead.
mugged .
23.3.3 .
makeae.
EASE .
EASE .
negate .
census .
meaetd.

EASE .

 

cem.omo
oom.cmo
oom.omo
cem.omo
oom.omo
cam.ome
osm.omc
oom.omc
oom.omo
oom.omc
cem.omo
oom.omc
cem.omo
oom.omc
oom.omo
cem.omc
oom.omc
oom.omc

com .omo

 

ohm.omm
Ohm.omm
ohm.omm
chm.omm
ohm.omm
chm.omm
chm.Omm
ohm.omm
ehm.omm
ohm.omm
chm.omm
chm.omm
Ohm.omm
chm.omm
ohm.omm
Ohm.Omm
chm.omm
ohm.Omm

cum 6?

 

mm—

mm—

mm—

mm—

mm

—

mm

—

mm

—

mm—

mm

—

_

mm

mm

mm

_

mm

_

mm

_

mm

mm

mm.

mm—

mm

_

.oom.mOm
.oom.wom
.oom.wom
.oom.woM
.oom.wOm
.oom.woM
.oom.wom
.oom.wOm
.oom.wom
.oom.wom
.oom.wom
.oom.wom
.oom.wom
.oom.wOm
.oom.mom
.oom.wom
.oom.woM
.oom.w0m

.oom .wom

 

omo

omo

omo

omo

omo

omo

omo

omo

 

EEO .Som
EEO Sow
EEO Sow
EEO Sow
EEO Sow
EEO .Som
EEO Sow
EEO Sow
EEO Sow
xtEO Sow
EEO Sow
EEO Sow
.CLEO Sow
htosOezom
EEO .Som
EEO Sow
EE O .Som
htosOeaom

CLEO Sow

 

wié

3%

3%

and

wmé

nmé

mmé

cmé

mmé

OHM

3A..

32m

awrm

xv-m

 

v—

E

3

v—

v—

3

v—

E

3

3

E

3

v—

v—

E

3

E

E

E

 

 

105

 

com

 

 

 

 

 

 

 

 

 

 

8.5% . - - .OON .OOO .OOO ONO Em. N55. ON
OON

83% . - - .OON .OOO .OOO ONO 2E .55. ON
OON

Seem . - - .OON .OOO .OOO ONO OS 5:5. .255. NN
OON

8.53... . - - .OON .OOO .OOO ONO OS 5.5 :55 NN

- - - - - - 252 .N

- - - - - - 252 ON

- - - - - - 252 o.

sees . OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 sow EOE O.

2328 . OOO .ONO ONO .OOO OO. .OON .OOO ONO 58,. .NNE N .

228.8 . OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 .35 .e . 6O. N.

2:28 . OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 25m :25 N.

miss . OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 .OOO E. :6 O.

3.28 . OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 .OOO w: .5 O.

sass . OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 .OOO E :6 O.

sass . OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 .OOO NNOEU O.

miss . OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 .OOO OOOO 3

 

 

 

 

 

 

 

 

 

106

 

 

 

 

 

 

 

 

88.8 O. OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 .OOO OENO 5 ON
88:8 .... OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 .OOO OON”. O... ON
8.8.... .w - - OO. .OON .OOO ONO €26 .OOO ONEO O... ON
88.8 .w OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 saw :3. O... ON
88.8 .... OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 .OOO NOSE ON
88.8 .O. OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 .OOO .ONNS ON
88.8 ..\ OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 5OO O: New ON
88.8 .4 OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 .OOO ENE ON
88.8 ..\ OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 .OOO NOONE NN
88.8 .w OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 .OOO .OONE NN
88.8 .... OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 .OOO NeOio ON
8.88 .4 OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 .OOO :25 ON
88.8 ..\ OOO .ONO ONO .OOO OO. .OON .OOO ONO €26 .OOO NeOio ON
- - - - - EEO Sow 2.02 E
OON
8.58w .w - - .OON .OOO .OOO ONO 88m 2%. ON
OON
838. ..O - - .OON OO.. .OOO ONO seem EOO. ON
OON
858. .w - - .OON .OOO .OOO ONO 88m O55. ON

 

 

 

 

 

 

 

 

 

107

 

 

63.536 .v.
EASE .v.
EASE .V
8263 .V
SSS: .V
6335 .V
32.58% .T
3326M .T
SEEM O.
SASS .V
3526M .Y
USES .T
SSS: .V
SSS: .v.
.OSEE .w
3553.. .v.
3556M .V

SASS .V

SSS: .v.

 

com .omc
cow 6%
Gen .08
0mm 6%
emm .owm

omm .owm

ocm .omo

cmm 6%
0mm 6%
9mm 6%

Omm 6%

com 58

omm . owm

 

ohm 6mm

Ohm 6mm

 

mmfl.oom.wom
mm_.oom.woM
mm_.oom.woM
mn_.chv
mn_.©hv
m>_.onv
mm_.com.woM
mm_.oom.wom
mm_.oom.wOM
mm_.oom.wom
mm_.oom.wom
mn_.cnv
mN_.ch
mb_.©nv
mh_.onv
mm_.oom.wom
mm_.oom.wom
mm_.oom.wom

m: .onv

 

omo

omo

omo

mum

m;

mum

omo

omo

cmc

mum

mum

mum

mum

omo

0N0

omo

mum

 

EEO Sow
EEO Sow
EEO Sow
EEO Sow
EEO Sow
EEO Sow
EEO .Som
EEO Sow
EEO Sow
EEO Sow
EEO Sow
EEO Sow
EEO Sow
EEO Sow
EOE O Sow
EEO .Som
EEO Sow
EEO Sow

EEO .Som

 

GEL—x y:

amt—m Mu—

owkod y:

_ __ hwy— M1—

NZBM m..—

mkhx y:

FEM y:

8.5. y:

Chm m1—

okom N:

cmbmx M1—

293. mu—

:sz y:

:an y:

0.,—hm mu—

ohmd y:

8.3— M:

trvm y:

8.3. y:

 

om

om

om

om

om

om

OON

om

 

 

108

 

 

EASE .
EASE .
EASE .
EASE .
EASE .
EASE .
EASE .
EASE .
EESE .
333M .
SEEM .
355mm .
EASE .
EEEEM .
EASE .
EASE .
EASE .

EASE .

 

ocm,omc
com.ome
oom.omc
oom.omc
oom.om©
oom.omo
com.omw
oom.omo

ocm .omo

com .26
com 6%
com .80

com 6%

 

chm.Omm
chm.omm
onm.owm
osm.omm
chm.omm
onm.omm
chm.cmm
own.omm

cum 63

Ohm 6mm

cum 63

05m 63

Ohm .omw

 

mm_.oom.woM
mm_.oom.wom
mm_.oom,woM
mm_.oom.w0m
mm_.ocm.w0m
mm_.oom.wom
mm_.oom.w0m
wm_.oom.w0m
mm_.oom.woM
ovm.mwm
mm_.oom.wOM

mm .oom .wom

m2 ,oom .wom
mm.— .coN .mOm
mm. .oom .wOm
mm. .oom .wom
m2 .oom ,woM

m2 ncom .wOM

 

omo

omo

omo

omo

omo

omo

omo

cmo

omo

omo

cmo

omo

omo

oma

omo

cmo

 

EEO Sow
EEO Sow
EEO Sow
EEO Sow
EEO .Som
PE O Sow
EEO Sow
EEO Sow
EEO Sow
EEO Sow
EEO Sow
EEO Sow
EEO Sow
b.05O Sow
EEO Sow
\EE O Sow
ztEO Sow

EEO Sow

 

mmhmm My.
mama mm
m 75% mm
:an mm
0.51 My.
”Ex My.
925m
mem MM
vhmx mm
nwhvm MM
emu—EM mm
2.5% MM
2.3— MM
mELM My.
”Ex My.
02% E M:—
mkim #3

33¢ y:

 

om

om

om

om

 

 

109

 

 

522: .V

355%

cSEmm .
355mm. .
353mm .
355mm .
3:3: .
3:2: .
333mm .
3:2: .
aEEmm .
3:22 .
magma .
3:2: .
8:2: .
makcad .
655mm .
uEEwm .

353mm .

.V

 

omm 6%

own 6%

omm ,owm

Omm ,owm

Omm dwm
com 6%
cmm .owm

omm .owm

 

cum 6%

 

ovm,cov
mm_.oom.wOm
mm_,oom.wOm
mm_.oom.wom
mm_.oom.wom
mm_.oom.wom

wn_.©nv
mh_.ovm,chv
mm_.oom,woM
mn_.cvm.cbv
mm_.oom.woM
mn_.ovm.©nv
wm_.oom.wom
mn_.ovm.chv
mn_.ovm.e>v
mm_.oom.woM
mm_.com.woM
mm_.oom.wom

m2 .oom ,woM

 

omo

omo

omo

omo

mum

mum

omo

mum

0N0

mww

omo

mum

mum

omo

omo

omo

 

€25 Sow
p.25 Sow
€25 Sow
Pogo .Som
90:0 Sow
ate—5 Sow
€25 Bow
p.25 .Som
€25 Sow
€20 Sow
€25 Sow
€25 Sow
€25 Sow
EEO Sow
@350 Sow
€25. Sow
$.25 Sow
€25 Sow

€25 50m

 

mhwmm mm

8va mm

Manama My.

Sigma My.

3:2? My.

3th MM

Zlhmmz My—

mmkomm my.

3.3:..— mm

wEbEyE

OPS—M my.

02%;— my.

753* My.

wgtx My.

59:; My.

2.5—y. My.

thwm My.

omlrwm My.

umhmm My.

 

om

om

om

om

om

am

am

om

om

om

om

om

om

om

 

 

110

 

 

33.53% .
33.53% .
33.53% .
33.53% .
33.53% .
33.53% .
33.53% .
33.53% .
33.53% .
33.53% .
33.53% .
33.53% .
3353% .
33.53% .
3.533 .
33.53% .
33.53% .

33.3.3333 .

 

com 6%

 

0:.

6mm

 

ovm.mww
ovm.mwm
o3~.mww
ovm,mwm
ovm.mwm
ovm.mwm
ovm.mwm
ovm.mwm
ovm.mww
ov~¢mwm
ovm.mwm
cvm.mwm
93m,mwm
ovm.mwm
03m,mww
ovm,mwm
03m.mwm

mm. .oom dam

 

oma

omo

omo

omo

omo

omo

omo

omo

omo

 

we; 5:3“.
we; 5:3“.
we; 23:3,.—
wog 5:3“.
wow. 5.23m
we; 5:3“—
woq cozmm
we; 5:3“—
woq 5:3“.
mo... 5:3,.—
woq 5:3“.
wou— 5:3“.
wou— cozam
m3 5:3“.
3332
5333m—
zooom

zuoom

 

o _ -oo-m-o_

o-©o-m-o_

w-co-m-o_

oboéflo _

m-©o-m-o_

36¢- m-o_

m-©o-m-o _

méo- m6 _

To???

In

mm

E

on—

mn—

~52

mhm

Cum

 

am

am

om

am

om

om

om

om

om

om

om

 

 

111

 

com

 

 

 

 

 

 

 

 

8.38.- . - - .28 .8.- .me cg we. 8:8“. 333 3m
8m

8.383 . - - .28. .28 .83 omo mo-. 8:8. a: 3
88.83 . 8m .83 - mm. .oom .wom o8 38:3 .83 ~58 mm
3.583 . 8m 5% - mm. .oom .33 88 €83 383m .Fm3m mm

- - - - - 920 .53m 3:32 3-
88.8 . 8m .88 EM 5% mm. 58 .32. 88 3.h. 8.8.... E; R
88.8 . 8m .83 Em 5?. mm. 58 gm o8 E23 sow EBB mm
888 . 8m .83 SM 53 mm. den .38 o8 92.3 .58 fine-3 mm
8.88 . 8m .83 EM 5% mm. 58 .38 omo b8.3 sow 3833 mm
8.88 . 8m .83 Sm 5?. mm. 58 .8». 8° 3.2.3 sow :85 mm
8385 . - - m: .38 E. E3... 8-8-2-3 8303 .m
8332 . - - m: .3: 8.. E23 sow 3-8-8-3 803 .m
8.88 . 8m .83 Em .on a. .28 .32 omo €23 883m Sm3m on
8.88 . 8m .83 EM 5% mm. 58 .o. m ca 3.2.3 82% .28 2
8.333 . - - SN .38 So 33 8:9,. 2-8-2: 8
8.383 . - - SN .38 ca 33 8...». . 78-2: 8

 

 

 

 

 

 

 

 

 

112

 

3.3... 3.. 3.3.3.3

3.3... .3 3.3.3.3

3.3... .3 3.2%.3

 

 

 

 

 

 

 

 

 

 

 

 

 

 

33.332 wEEB 33.33,.- 332333 33.3%32 wEEB 33.333- wEEE 353-r flam—
._3... .3 3.3.3.3 3.3.... 33 3.3.3.3 3.3... 3.. 3.3.3.3

33.332 @335. 33.33% 33:83 33832 wEEB 33.33% wEEB 33.333- MID—m—
tg... 3.. 3.3.3.3 :3... 3.. 3.3.3.3 3.3... .3 3.3.3.3

3>_.3w32 wEEB 3833-3 32:83 3>_.3w32 wEEB 33.333. wEE... 33.33% maim—
._3... 3 3.3.3.3 3.3... .3 3.3.3.3 3.3... .3 3.3.3.3

33.33.32 wEEB 33.33..- 3>Emom 33.3%32 wEEB 33.333- wEEE 33.333- :35.
3.3... .3 3.3.3.3 3.3... .3 3.3.3.3 3.3... .3 3.3.3.3

33.3%32 mic-.2 3333-.- 3>Emom 33.3332 wEEE 388,—- wEEs. 33.33..- 3.52m
3.3... .3 3.3.3.3 3.3... .3 3.3.3.3 3.3... .3 3.3.3.3

3>_.3w3 Z wEEB 33.333- 3>Emom 33.33.32 wEES 33.33,.- wEEB 33.333- EmIm
3.3... .3 3.3.3.3 3.3... 33 3.3.3.3 :3... .3 3.3.3.3

3>_.3w3 Z wEEB 3833-.- 3>Emom 33.3%32 wEEB .233- wEEE 33.83- _ 3. _ 43
:3... .3 3.3.3.3 3.3... .3 3.3.3.3 3.3... .3 3.3.3.3

32.33.32 wEEB 33.3-33- 33333 33.3.32 3.3.3.... 32333- wEEB 33.333- Two-mm-c-Bm
3.3... .3 3.3.3.3 3.33.. .3 3.3.3.3 3.33.. 3.. 3.3.3.3

33832 9:53 32333- 33533 32.332 93...... 3833-.- wEEB 33.33% m-mo-o _ 6-3m
3.3.... .3 3.3.3.3 3.3... .3 3.3.3.3 3.3... 33 3.3.3.3

3533 Z wEEB 33.33% 33:83 3.5332 wEEB 33.33,.- wEEB 33.33% Vmo-w-o-Bm
3.3... .3 3.3.3.3 3.3... .3 3.3.3.3 3.3... .3 3.3.3.3

33.33.32 wEEB 33.33% 32:83 32.332 @353 33.333- wEES 33.33% m-mo-w-c-Bm
:3... .3 3.3.3.3 3.3... 3.. 3.3.3.3 :3... .3 3.3.3.3

33.3%32 wEEB 33.33% 32:33.. 33332 wEE... 33.83- %EEE 38.333- Ymo-mé-Bm
=3... .3 3.3.3.3 333.. .3 3.3.3.3 3.33.. .3 3.3.3.3

32.33.32 wEEB 33.333. 33:83 33.33.32 wEEB 33.33% @353 33.333 Two-m-o-Bm

.._> 3.22. 72. .._> 8.32. ::.> £22. .3... 33.32. c: 33.32. c: 8.32.
3353.: o _ .33. 3353... 363 3353.3 m- _ m 3353... 3.-. _ 3353... T. _ 3353:.
353.235.3353. 353235.535 353235.355 353.235.32.33 353235.32... 353.235.3253. 3.3.3.3—

 

 

 

 

 

 

 

 

333...— mdmm .35. .833.— TmU— 2: .33

5.335.353 m0.— wEm: 33.53% .V 53333333533 .V 3.. 635.53 35.335.35.3V33 353-cm 38 3.333.. 3.3.3.3—.9533 _a=x3m .~-< 33.:-

113

2.383... 3.3:. :33... 33.... 33.3.33. .33. 5.3 33.333283 3... 335333 6-. _ 3:3 7: 333.33. 5.3 33.332333 3.33.3 33.33..- .1.
33.333 3833 3... mo 33.3.33. 33.33. 5.3 33333.3 33:3 :35 E33,. 3.33:. 3;. o. 33.332333 33>3= 3.3.33.3 *

 

33.332

:3... 3.. 3.3.3.3
wEFS. 33.33,.-

32.333

33.332

3.3.3.4.. 3.3.3.3
wEE... 33.333-

...3... .3 3.3.3.3
wEEB 33.33%

3. OH

 

33.3%3 Z

3.3... 3.. 3.3.3.3
wEE... 33.33%

33.2333

32.3%32

3.3.... .3 3.3.3.3
wEE... 33.33%

3.3... .3 3.3.3.3
wEE... 33.33%

W-D.—-

 

3>_.3w3z

3.3.... .3 3.3.3.3
wEEs. 33.333-

3.5.8..

3>_.3w3z

3.3... 3.. 3.3.3.3
wEE... 33.33%

3.3... .3 3.3.3.3
9:5... 3.333-

O _ woo-.92

 

33.3%32

3.3.... .3 3.3.3.3
93...... 33.333-

32:33..

33.3%32

3.3... .3 3.3.3.3
wEE... 33.33..-

3.3... .3 3.3.3.3
wEEs. 33.33%

E

 

33.332

3.33.. .3 3.3.3.3
wEE... 33.333-

33:33..

3>_.3w3z

3.3.... .3 3.3.3.3
@353. 33.333-

..3... .3 3.3.3.3
mic-S. 33.333-

nmhvm MM

 

 

33.332

 

:3... 3.. 3.3.3.3
wEE... 33.333-

 

33:33..

 

3>..3w3z

 

3.3... 3.. 3.3.3.3
MEE... 33.33,.-

 

..3... 3.. 3.3.3.3
@353. 33.33%

 

92m—

 

 

114

 

 

 

 

 

 

 

 

 

 

 

 

 

 

3>..3m37. 9:5... 33.33.. 3.0.3.3 33.33 F 3.0.3.3 3.33% 3>...303 w=.E=. 33.33 .3 mm..wy. y.y.
3.33.. 3.. 3.3.3.3 3.33.. 3.. 3.3.3.3

3>..3w3Z wEE... 33.33.. 3.0.3.3 33.33.. 3.0.3.3 33.33 .. 3>...303 w=.E:. 33.33 .. ”Fwy. y.y.
3.33.. 3.. 3.3.3.3 3.33.. 3.. 3.3.3.3

3>..3w32 wEE... 33.33% 3.0.3.3 33.33 .3 3.0.3.3 33.33 P 3>...303 wEE... 33.33.. «hwy. y.y.

3>...303

3.33.. 3.. 3.3.3.3 3.33.. 3.. 3.3.3.3

3>..3w3Z wEEB 33.33.. 3.0.3.3 .23.. 3.0.3.3 33.33.. 32.33.32 wEE... 33.33.. m.....y. y...
3.33.. 3.. 3.3.3.3 3.33.. 3.. 3.3.3.3

3>..3w3z 3.3.333. 33.33.. 3.0.3.3 33.33 .. 3.0.3.3 33.33.. 3>...303 wEE... 33.33 .3 am: .y. y...
3.33.. 3.. 3.3.3.3 3.33.. 3.. 3.3.3.3

3>...303 9:35. 33.33 3 3.0.3.3 3.33.. 3.0.3.3 33.33 .. 3>..3w3Z w:.E=. 33.33 h 335. y...
3.33.. 3.. 3.3.3.3 3.33.. 3.. 3.3.3.3

3>..3w3Z wEE... 33.33.. 3.0.3.3 33.33 .3 3.0.3.3 33.33 h 3>...303 w=.E=. 33.33.. PSy. y...
3.33.. 3.. 3.3....3 3.33.. 3.. 3.3.3.3

3>...303 wEEB 33.33 P 3.0.3.3 33.33.. 3.0.3.3 33.33.. 3>..3w3Z wEEB 33.33.. 32.3. y...
3.33.. 3.. 3.3.3.3 3.33.. 3.. 3.3.3.3

3>...303 wEE... 33.33.. 3.0.3.3 33.33 H 3.0.3.3 33.33 .r 3.332 wEE... 33.33 3 3..wy. y...
.33.. 3.. 3.3.3.3 3.33.33 3.3.3.3

3>...303 wEE... 33.33 P 3.0.3.3 33.33.. 3.0.3.3 33.33 .. 3>..3w3z wEEB 33.33.. 3.33. y...
3.33.. 3.. 3.3.3.3 3.33.. 3.. 3.3.3.3

3>..3w3Z wEE... 33.33 .. 3.0.3.3 33.33.. 3.0.3.3 33.33 .. 3>...303 wEEa. 33.33 .. 3 . ..Ny. y...
.33.. 3.. 3.3.3.3 3.33.. 3.. 3.3.3.3

3>..3w3Z wEEz. 33.33.. 3.0.3.3 33.33 .r 3.0.3.3 33.33 .3 32.303 333.333. .3.33.. . .. . y. y...

... 3322. 3-33. 2. 33.32. 3 33.3.2. 3 33.32. .. 33.3.2. 3 333.2.
333.3... .733. 333.3... 3-3 33.33... :3. 33333... 34.3. 333.33... 3.3 333.3...
33.3..3333..E3.3 3.33.33.32.23 3.33.33.35.33. 33.3.3333...E.3 33.33.333.35... 3.33.33.33.32... 3.3.03.

 

 

 

 

 

 

 

 

3:33.33 many. 3:3 30.33 ..m6— 3...

.0 3.335.353 m0.— w:.3= 33.53% .33 33.33.0330 .33 33 33......33. 3.55.353V? 3533.3 30. 3....333 b.......33503 .asxom .m-< 333,—.

115

 

 

 

 

 

:3: B 22% :8: 3 22%

02:8; wEEB Bach 229v .Bmoh ESE—u $58. ozumwoz wEES Buck mFmUm
:3: B 22% :3: B 22%

328m wEEB 533‘ 2036 5.8% 203% 5.8% 03832 wEEE .889 _ hmom
(:3: B 22% :3: 322%

02832 wEEE $38K ROBE Bach 229v Sufi. 02:8; wEEa 57.0% _ .5 _ m ”E
(:3: E 22% :3: E 22%.

oZSwoZ wEEE Back 2036 Back 2293 .889 358m wEEB .889 ovho _ x mm
:8: 3 22% :3: B 22%

ozawoz wEEB 5.8% 203% Back 22a? 528% 328m wEEB .38? o _ .2: x mm

 

 

 

:3: E 22%

 

 

 

 

:3: 3 22%

 

 

 

116

APPENDIX B:

SOMATIC INCOMPATIBILITY TESTS AND MICROSATELLITE RESULTS

Table B-1: Results from somatic incompatibility test and microsatellites for strains
of A. ostoyae collected from orchard A (site # l4) and orchard B (site # 29) and
microsatellite results for six A. ostoyae isolates and one A. gemina from

miscellaneous sites

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Isolate Site SIG CAG25 CAG77 AOSSR74 AOSSR84
1-59 l: 0511-1 356 327 126/137 92/94/96
1-101 14 0511-1 356 327 126/137 92/94/96
1-217 14 0511-1 - - - -

2-27 14 0512-1 353 322/324 123/137 98/100
243 14 0512-1 353 322/324 123/137 98/100
244 14 0512-1 - - - -
2-48 14 0513-1 353/356 322/324 135/137 96/100
2-49 14 0513-1 - - - -
2-91 14 0513-1 353/356 322/324 135/137 96/100
3-64 14 0514-1 348/356 327/336 126/130/135/137 90/96
3-79 14 0514-1 - - - -

4-280 14 0514-1 - - - -

4-306 14 0514-1 348,/356 327/336 126/130/135/137 90/96
4-35 14 0515-1 353 327 128/130/132/135 82/92/94/96/98
4-55 14 0515-1 - - - -

4-57 14 0515-1 353 327 128/130/132/135 82/92/94/96/98
4-82 14 0515-1 - - - -
4-148 14 0515-1 - - - -

 

 

 

 

 

 

 

117

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

4-36 14 0516-1 348/356 330 126/130/137 92/94/96/98
4-58 14 Ost6-l - - - -
4-59 14 0516-1 - - - -
4-86 14 0516-1 348/356 330 126/130/137 92/94/96/98
LR RlTl 29 0511-2 351/353 327/330 122/137 82/86/92/104/10
6
LR R2T9 29 0512-2 - - - -
LR R417 29 0512-2 356 336 122/126/128/130 86/90/94
LR R6TO 29 0512-2 - - - -
Fl 29 0512-2 - - - -
LR R21“! 9 29 0513-2 356 327/3 36 122/1 28 94/96/98/100
LR R9T80 29 0514-2 - - - -
LR R1 ”‘38 29 0514-2 - - - -
LR R11T4o 29 0514-2 - - - -
LR Rl5T39 29 0514-2 351 322/332 128/132 90/92/94/98/104
LR R 14T4 29 0515-2 - - - -
LR R14T5 29 0515-2 348/356 327 126/128 86/92/94/98
RR RIT8 29 0516-2 - - - -
RR R5T4 29 Ost6-2 - - - -
RR R6T2 29 0516-2 353/356 330/3 36 132/137 82/88/92
RR R7Tl 29 0516-2 - - - -
RR R8T15 29 0516-2 - - - -
RR R4T3 29 0517-2 351 332 128/1 30 88/90/92/94/98
RR R10T19 29 0517-2 - - - -
RR R5T8 29 0513-2 - - - -
Stump 29 0513-2 353/356 332 132/135/137 92/100/102/104/
l06

 

118

 

 

RR R6T6 29 Ost9-2 - - - -

 

RR R8T22 29 Ost9-2 - - - -

 

 

 

 

RR R8T23 29 Ost9-2 351/353 330 132 92

RR R10T40 29 Osth-l 351/353 327 126/128/135/137 82/92/98/102

RR R18Tl 29 Ostl 1-1 351/356 327/336 128/137 90/94/100/102/1
04

LR R1T73 29 Geml-2 348/351 330 126/128 90/92/94/96

 

RR R1T73 29 Geml-2 - - - -

 

RR R4T35 29 Geml-2 - - - -

 

RR R4T36 29 Geml-2 - - - ..

 

RR R8T34 29 Geml-2 - - - -

 

RR R8T36 29 Geml-2 - - - -

 

RR R8T42 29 Gem1-2 - - - ..

 

RR R18T50 29 Geml-Z - - - -

 

RR R19T45 29 Gem l-2 - - - -

 

RR R23T62 29 Geml-Z - - - -

 

RR R23T63 29 Geml-2 - - - -

 

RR R24T72 29 Geml-Z - - - -

 

RR R24T73 29 Geml-2 - - - -

 

RR R24T97 29 Geml-2 - - - -

 

LR R5T6 29 Gem2-2 348 330/346 128/130/132 90/92/94/96/ 100

 

LR R5T9 29 Gem3-2 - - - -

 

LR R617 29 Gem3-2 351 330/346 128/130/132 90/92/94/96/104

 

LR R7T6 29 Gem3-2 - - - -

 

LR R7T7 29 Gem3-2 - - - -

 

LR R5T56 29 Gem4-2 348 330/346 128/130/132 90/92/94

 

 

LR R4T6 29 Mell-2 - - - -

 

 

 

 

 

 

 

119

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

LR R5T10 29 Mell-2 -
LR R5T1 l 29 Mell-2 -
LR R5T15 29 Me11-2 -
LR R5Tl6 29 Mel 1-2 -
LR R7T8 29 Mell-2 -
LR R7T12 29 Mell-Z -
LR R8Tl 1 29 Mel] -2 -
RR R17T27 29 Me12-2 -
RR R17T28 29 Mel2-2 -
RR R18T29 29 Me12-2 -
RR R19Tl8 29 Me12-2 -
RR R20T25 29 Me12-2 -
RR 29 Mel3-2 -
R22T141
RR R28T3 29 Mel4-3 -
RR R4T57 29 Gal 1-2 -
BTl 29 Gal2-2 -
BT3 29 Ga12-2 -
MT2 29 Ga12-2 -
F5 29 0312-2 -
F6 29 Gal2-2 -
F7 29 (3312-2 -
F8 29 GalZ-2 -
F11 29 Gal2-2 -
10-3 -06-1 29 Gal3-2 -
10-3 -06-2 29 GaI3—2 -
10-3-06-3 29 Gal3-2 -

 

 

 

 

 

120

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

10-3-06-4 29 Gal3-2 - - - -
10-3-06-5 29 Gal3—2 - - - -
10-3-06-6 29 Gal3-2 - - - -
10-3-06-8 29 Gal3-2 - - - -
10-3-06-9 29 Gal3-2 - - - -
10-3-06-10 29 Gal3-2 - - - -
10-3-06-1 1 29 0313-2 - - - -
l 0-3-06- 12 29 6313-2 - - - -
PT1T2 3 - 342/356 330/336 124/130/132/134 94/98/100
CT12T1 l7 - 344/356 330 126/128/130 90/94/100/102
CT 1 9T2 26 - 356 322/327 124/128 80/84/86/90/92/
102
SC2T1 3O - 356 322/327 130/132 82/92/94/96/98
CT23T1 32 - 346/356 330 126/132 84/88/90/102
XTlTl 33 - 353/356 327/330 132/134 84/92/98/100
SC3Tl 35 - 344/348 327/330 120/122/124/126/128 80/90/92/94/100

 

 

 

 

 

 

 

121

 

APPENDIX C:

GREENHOUSE ROOTSTOCK INOCULATION AND FIELD TRIAL

INTRODUCTION

Many orchard sites in Michigan are being converted into vineyards because
grapes have more value per acre. However, growers are often unaware that Armillaria
has been reported on grapes in Australia, Brazil, Central and Eastern Europe, and in
California. Even if a grower is aware of the potential problem, new growers may not
know the history of an orchard and whether or not Armillaria is present. Michigan’s
grape industry is much younger than those of both Europe and California and the effects
or presence of infection with A rmz'llaria have not yet been reported. Michigan does have
one of the species of Armillaria reported on grapes, A. mellea, and another potentially
pathogenic species, A. ostoyae. In the Northwestern cherry producing region of
Michigan where many of the vineyards are currently being established, A. ostoyae tends
to be the predominant species and it is known as a virulent pathogen with a wide host
range. While most reports of A. ostoyae are on conifers, it is proven to be an aggressive
pathogen of cherry in Northwest Michigan, as seen by Proffer et al. in the 1987 survey
and rootstock inoculation trials (Proffer et al. 1998). Its continued impact on cherry was
confirmed by the current survey.

In California, Baumgartner et al. (2006ab) have begun to test grape rootstock
susceptibility to A. mellea. They tested eight different rootstocks finding rootstocks
3309C and Riparia Gloire, commonly used in Michigan, most susceptible to A. mellea,

while Freedom, not commonly used in Michigan, was found to be most resistant. Other

122

rootstocks that commonly do well in Michigan soils were not tested by Baumgartner,
including S-BB, s04, 101-14 Mgt (Howell et al. 1999). With the increase of viticulture '
in the state of Michigan, it is important to begin and assess A. ostoyae and A. mellea for
their possible pathogenicity to grape rootstocks grown in Michigan. The hypothesis of
this work was that the current Armillaria species known to infecting orchard grown
Prunus in Michigan will move to grape when orchards are converted to vineyards. The
hypothesis was tested by beginning greenhouse and field trial inoculations of grape
rootstocks grown in Michigan for the future assessment of susceptibility to A. ostoyae

and A. mellea.

MATERIALS AND METHODS: GRAPE ROOTSTOCK SELECTION

Four varieties of grape rootstocks, 50 rootstocks per variety, were ordered from
California Grapevine Nursery Inc. (St. Helen, CA 94574-9790). The grape rootstocks
included the following varieties: Freedom, Riparia Gloire, 3309C, and 101-14. Although
not used in Michigan, Freedom was chosen as a negative control since it was previously
shown to be the most resistant rootstock to A. mellea by Baumgartner et al. (2006ab).
The positive controls included Riperia Glorie and 3309C. They were selected because
they are commonly used in Michigan, and were known to be susceptible to A. mellea
(Baumgartner et a1 2006ab). The rootstock 101-14 was chosen because it is very
commonly used in Michigan. Once received, the rootstocks were temporarily stored in a
cold room until planting to prevent budding. The rootstocks were then brought to room

temperature (20° C to 25° C) four days before planting.

123

INOCULUM PREPERATION

Inoculum was prepared three months prior to planting. To prepare the inoculum
Prunus mahaleb twigs approximately 2.5 cm in diameter were cut from spot trees
near-an old orchard site at Michigan State University. The twigs were cut into
approximately 8 cm pieces and were then autoclaved in a bag with 100mL water for
30 minutes. After being autoclaved 25 twigs per bag were placed into 16
autoclavable fungal spawn bags containing. These bags have an air vent and are sold
commercially for culturing edible mushrooms. In addition to the twigs, 500mL of 4%
MBA amended with 20 g/l glucose and 5 g/l peptone was added to each bag. The
tops of the bags were folded and closed with autoclave tape, and autoclaved for 30
minutes. Once autoclaved the bags with the media and twigs were allowed to cool
until the media was partially solid and 400 uL of 90% ethanol were added to the bag
under aseptic conditions. The bags were inoculated with three 60 mm by 15 mm
Pertri dishes of one isolate of Armillaria (Table C-l for isolates of Armillaria used).
For each strain there were a total of two bags. The Armillaria isolates used were
primarily collected from Michigan orchards. One isolate of A. mellea was not a
Michigan isolate, since at the time there were not enough strains of this species in the
collection at the time of inoculation. The inoculated spawn bags were then sealed
using a seal-a-meal machine and placed in the dark at 25° C for three months (Figure

G] for prepared inoculum).

124

Table 0]: Strains of Armillaria used to inoculate grape rootstock

 

 

 

 

 

Strains of Armillaria
A mellea isolates: A ostoyae isolates:
1. B277 UNH Harrington l. PT1T2
2. C2 23-2 x 10-1 2. CT9T1
3. 1-1x11-20 Cl 3. CT20T1
4. CT1T2 4. DC. Co. 5-19-05-1
GREENHOUSE SETUP

The 160 rootstocks, 40 of each variety, were planted in 3 gallon pots filled with
one part sand and one part soil obtained from the greenhouse at Michigan State
University. The rootstocks were then inoculated on May 19, 2006 with three different
isolates of either A. ostoyae or A. mellea (Table C-2 for setup). There were four
replicates for each inoculum combination. The inoculum pieces were placed next to the
rootstock and were buried approximately 2.5 cm into the soil mix. A layer of mulch was
then placed on top of the inoculated pot to keep the soil cool. The inoculated grape
rootstocks were placed outside for the summer (Figure C-2). The pots were then moved
into the greenhouse for the winter at the end of September and were again moved outside
in May of the following spring.

The plants were checked on regular bases for signs of decline, such as defoliation.
At nine months every rootstock was thoroughly checked for signs of mycelial fans under
the bark of the roots, and also for signs of rhizomorph growth from the inoculum source.
Each inoculum source was also checked to make sure it was still viable. All pots
inoculated with A. mellea still had viable inoculum, however most of the A. ostoyae pots
no longer had viable inoculum. All of the A. ostoyae pots were re-inoculated with the

PTlTl strain.

125

Table 02: Pot setup for greenhouse inoculations for each rootstock variety

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

A. ostoyae A. mellea A. ostoyae A. mellea
Rep isolates Rep isolates Rep isolates Rep isolates

1 1,2,3 1 1,2,3 3 1,2,3 3 1,2,3
1 2,3,4 1 2,3,4 3 2,3,4 3 2,3,4
1 3,4,1 1 3,4,1 3 3,4,1 3 3,4,1
1 4,1,2 1 4,1,2 3 4,1,2 3 4,1,2
1 CK 1 CK 3 CK 3 CK

2 1,2,3 2 1,2,3 4 1,2,3 4 1,2,3
2 2,3,4 2 2,3,4 4 2,3,4 4 2,3,4
2 3,4,1 2 3,4,1 4 3,4,1 4 3,4,1
2 4,1,2 2 4,1,2 4 4,1,2 4 4,1,2
2 CK 2 CK 4 CK 4 CK

FIELD TRIAL SET UP

' The four grape rootstocks were also used for a long term field trial to see if
Armillaria ostoyae will move to grape in the field. The site chosen for the field trial was
a peach orchard in Leelanau County where A. ostoyae was infecting peach trees. The
growers were interested in converting the orchard to a vineyard, making this site ideal for
a rootstock test. The results will give the grower, as well others, valuable information on
the suitability of Armillaria sites for vineyard production. Ten rootstocks of each type
were planted on May 16, 2006. The rootstocks were planted within an infected pocket
with root debris and stumps with viable Armillaria infections present. Each of the

rootstocks was planted 1 m apart in 2 rows (Figures C-3 and C-4).

126

 

 

FIGURE C-l
Prepared Armillaria inoculum for the greenhouse inoculation trial to test grape

rootstocks to susceptibility to A. ostoyae and A. mellea

 

Fig C-l: Inoculum prepared using cherry twigs for greenhouse inoculation to test grape
rootstocks for susceptibility to Armillaria.

127

FIGURE C-2:

Greenhouse inoculation trial to test four different grape rootstocks to their

susceptibility to A. ostoyae and A. mellea

 

Fig C-2: Grape rootstocks inoculated in a greenhouse trial with Armillaria ostoyae and Armillaria mellea
strains.

128

FIGURE C-3:

Grape rootstock field trial to test four rootstocks to susceptibility of A. ostoyae

 

Fig C-3: Grape rootstock planted in an infected peach orchard to test for susceptibility to A ostoyae. Grape
rootstocks are surrounded by a layer of much and are planted in two row going out from a central source
towards two other sources.

129

FIGURE C-4:

Field trial setup to test grape rootstock to susceptibility to A. ostoyae

» , Rootstock Variety 1 . Rootstock Variety 3 g

Rootstock Variety 2 Rootstock Variety 4

 

130

CONCLUSIONS

To date there have been no signs of infection due to Armillaria from either the
greenhouse or field trial. Symptoms caused by infection from Armillaria root rot should
not be expected for a couple of more years in the field trial based on the rate at which
symptoms are seen in tart cherry from field observations. The rootstocks from the
greenhouse trial will be check at the end of August to see if any symptoms have
developed. If Armillaria spp. found in Michigan are able to infect grape there will
probably be more success with the field trial where syptoms should be seen if the
rootstock are infected. In greenhouse inoculation trials, Baumgartner (2006a,b) reported
that although infection occurred, no symptoms, such as defoliation, occurred even 12

months after inoculation.

131

REFERNCES

Baumgartner, K. & Rizzo, D.M. (2006a). Relative resistance of grape vine stock to
Armillaria root disease. American Journal of Enology and Viticulture. 7: 408-414.

Baumgartner K. & Rizzo, D. M. (2006b). Resistant grapevine rootstocks for control of
Armillaria root disease. American Journal of Enology and Viticulture. 57: 383A-
383A.

Howell, G.S., Miller, D.P., & Zabadal, T.J. (November 1999). Wine grape varieties for
Michigan. Retrieved July 3, 2006 from the Michigan State University Extension
webpage: httpz/lwebl .msue.edu/ imp/modfr/2643 9701 .html.

Proffer, T. J ., Jones, A.L., & Ehret, GR. (1987). Biological species of Armillaria isolated
from sour cherry orchards in Michigan. Phytopathology. 77: 941-943.

Proffer, T. J ., Jones, A.L., & Perry, R.L. (1988). Testing of cherry rootstocks for
resistance to infection by species of Armillaria. Plant Disease. 72: 488-490.

132

APPENDIX D:
GLOSSARY

GLOSSARY:
Acropetal: Proceeding from the base toward the apex or from below upward, used to
describe the movement of fungicides in a plants, so in this case it would move into the
stem or trunk of a tree.
Adnexed: Pertaining to the attachment of the fertile tissue (the gills, tubes, spines, etc.) to
the stipe of the fungus in which the fertile tissue typically curves upwards towards the
pileus of the fungus before attaching to the stipe.
Arachnoid: See Cortina
Basidiome: Mushroom, basidiocarp or fruiting body that bears basidiospores.
Basipetal: Proceeding from apex towards the base or from above downward, used to
describe the movement of fungicides in a plants, so in this case it would move into the
root system.

Binding hyphae: Thick-walled, typically aseptate, highly branched vegetative hyphae.

Caespitose: When basidiomes grow in dense clusters, with the stems fused together or
packed right up against one another at the base.

Cortina (Cortinaceous): A special type of annulus that is filamentous, resembling a spider
web, attached from the margin of the cap (pileus) to the stem (stipe) when young. In age
only a few fibers may remain on the cap margin or the stipe.

Clamp connection: A bridge-like hyphal connection involved in maintaining the
dikaryotic condition.

Decurrent: Pertaining to the attachment of the gills to the stipe, in which the gills curve
partly down the stipe towards the base of the stipe.

Dichotomous: A type of hyphal branching into two equal forks.
Dikaryon: A hypha or portion of hyphae which contains two haploid nuclei in each cell.

Floccose: Having a cottony appearance, like a flocked Christmas tree. Seen in some
species of Amanita.

Generative hyphae: Undifferentiated, thin-walled, usually have frequent septa, and may
or may not have clamp connections.

133

Gregarious cluster: When basidiomes grow in a group that is a bit more scattered and
irregular.

Karyogamy: Fusion of two nuclei.

Monokaryon: A hypha or portion of hyphae which contains one haploid nucleus in each
cell.

Monopodial: A type of hyphal branching which maintains a single direction of growth
and splits laterally.

Plasmogamy F
Primary mycelium: Mycelium prior to dikaryotization, produced by basidiospores.

Pseudosclerotial plate: See zone line.

 

Secondary mycelium: Mycelium subsequent to dikaryotization, or the fusion of primary .'
mycelium from two different mating types. ll

Skeletal hyphae: Thick-walled and very long and straight, with little cell content. They
have few septa and lack clamp connections.

Sinuate: Referring to a type of gill attachment, specifically gills that are notched at their
point of attachment to the stipe.

Subhymenium: Refers to the layer of cells between the hymenium, the layer of spore-
bearing cells, and the main trama of the gills, or tubes, of a basidiome.

Zone line: black lines seen in decayed wood, which are curved planes in the wood,
composed of thickend, dark fungal cells

134

11111111111111111111111111