Molecular Characterization of Southeastern Armillaria Isolates

MOLECULAR CHARACTERIZATION OF SOUTHEASTERN ARMILLARIA ISOLATES

KAROL LEIGH KELLY

(Under the Direction of Kathryn C. Taylor)

ABSTRACT

The genus Armillaria contains white-rot basidiomycetes pathogenic to woody plant hosts worldwide. Of particular interest are the species found in southeastern peach (Prunus persica) orchards causing Armillaria root rot. Isolates collected during the past four years from orchards in Georgia, South Carolina, and Alabama were identified to species and grouped within species based on molecular analysis of the internal transcribed spacer (ITS) regions and the intergenic spacer (IGS) regions of the ribosomal DNA (rDNA). Thirty isolates of A. tabescens and nine

isolates of A. mellea were identified from these orchards. Restriction fragment length

polymorphism (RFLP) analysis of the IGS region with Alu I revealed two groups of A. tabescens

and one group of A. mellea. Similarly, the ITS region was amplified with primer sets At-

ITS1/Am-ITS1/ITS-2 and ITS-1/ITS-4 and subsequently restricted with Mbo II and Hha I.

These restrictions indicated two groups of A. tabescens and four groups of A. mellea.

INDEX WORDS: Armillaria mellea, Armillaria tabescens, Armillaria root rot, Basidiomycetes, intergenic spacer region, internal transcribed spacer region, peach, polymorphism, Prunus persica, rDNA, restriction analysis

MOLECULAR CHARACTERIZATION OF SOUTHEASTERN ARMILLARIA ISOLATES

KAROL LEIGH KELLY

B.S., Georgia College and State University, 1996

A Thesis Submitted to the Graduate Faculty of The University of Georgia in Partial Fulfillment

of the Requirements for the Degree

MASTER OF SCIENCE

ATHENS, GEORGIA

2004

Karol Leigh Kelly

MOLECULAR CHARACTERIZATION OF SOUTHEASTERN ARMILLARIA ISOLATES

KAROL LEIGH KELLY

Major Professor: Kathryn Taylor

Committee: Mark Rieger Harald Scherm

Electronic Version Approved:

Maureen Grasso Dean of the Graduate School The University of Georgia August 2004

DEDICATION

I would like to dedicate this thesis to my wonderful family. My parents, Sam and Judy

Kelly, have believed in me since the very beginning. They are the most humble, honest, and wonderful people that I know. I am so lucky to have them in me life. They have taught me that I can do anything that I put my mind to and that I should always look for the best in people. I don’t think there are better parents anywhere. I love y’all with all of my heart.

I would also like to dedicate this to my sister Lisa. She is good at keeping me honest in everything that I do. She also has the best heart of anyone I know and will make a positive difference in the world. I love you.

Finally, I would like to dedicate this to my split-apart, Kristen. There is not a doubt in my mind that I would not have made it through this research without your help. You have been my cheerleader, my shoulder to cry on, and my best friend through it all. I look forward to spending the rest of my life with you… I love you.

ACKNOWLEDGEMENTS

Not only do I have the most wonderful family in the world, I think I also have the best

friends. Everyone was such a help to me whether it was as a mentor, proof-reader, or the most

important of all moral support – most involved had multiple rolls. I couldn’t have undertaken

this work at all without the support of Kathy Taylor. I’ve been fortunate to have you as a boss,

mentor, professor, and pain in my butt (I meant to say friend). I never would have had the

opportunity for this degree had it not been for you.

Stacy, maybe one day you will be able to see the river despite the forest (that you tried to

drive through). Thanks for all that you have done for me. You are a wonderful friend and huge

pain in the butt. Janice and Merry (and Charlotte and Laura), I couldn’t have made it without all of your therapy sessions and proofing. Y’all have been good friends and support to me. Carol and Henry, you were a continuous source of wisdom, inspiration, and laughs. I am so thankful that those random chance meetings have made us friends. I am continually awed by your intellect and kindness – not many people have such a wonderful combination.

Dr. Scherm, Dr. Rieger, and Dr. Beckman, I know that I could not have made it without your persistent, driving forces and advice. I am so lucky to have had y’all as mentors and instructors during this time. I have learned so much from you. I count myself lucky to have been your student.

TABLE OF CONTENTS

Page

ACKNOWLEDGEMENTS...... v

LIST OF TABLES...... vii

LIST OF FIGURES ...... ix

CHAPTER

1 INTRODUCTION ...... 1

2 MATERIALS AND METHODS...... 20

3 RESULTS ...... 31

4 DISCUSSION...... 43

REFERENCES ...... 55

LIST OF TABLES

Page

Table 1: Armillaria isolates collected from southeastern peach orchards and used in the current

study...... 28

Table 2: Polymerase chain reaction (PCR) primers used to amplify regions of Armillaria

DNA…...... 29

Table 3: Summary of the molecular analysis of known and unknown Armillaria isolates..... 35-36

Table 4: Identification and classification of southeastern Armillaria tabescens isolates...... 52

Table 5: Identification and classification of southeastern Armillaria mellea isolates...... 53

vii

LIST OF FIGURES

Page

Figure 1: Schematic illustration of the typical arrangement of rDNA of basidiomycetes,

including primer locations...... 19

Figure 2: Geographical distribution of Armillaria isolates collected from southeastern peach

orchards ...... 30

Figure 3: Analysis of the IGS and ITS of known A. tabescens isolate TA1 and known A. mellea

isolate ST5 ...... 37

Figure 4: IGS region of Armillaria isolates amplified with primers LR12R/O-1 and restricted

with Alu I...... 38

Figure 5: IGS region of Armillaria isolates amplified with primers based on the sequences of

European isolates (Sicoli et al., 2003) to identify A. mellea and A. tabescens…...... 39

Figure 6: ITS region of Armillaria isolates amplified with primer set ITS-1/ITS-4 and the primer

set developed by Bryson et al. (2003) to discriminate among southeastern Armillaria

spp...... 40

Figure 7: ITS regions of putative A. tabescens isolates amplified with primer set ITS-1/ITS-4 and

restricted with Hha I and Mbo II to visualize polymorphisms among isolates...... 41

Figure 8: ITS regions of putative A. mellea isolates amplified with primer set ITS-1/ITS-4 and

restricted with Hha I...... 42

Figure 9: Geographical distribution of Armillaria “groups” as established by molecular

characterization of IGS and ITS region...... 54

viii CHAPTER 1

INTRODUCTION

Purpose of the Study

The southeastern peach industry. According to the National Agricultural Statistics

Service (NASS-USDA, 2004), 45,350 metric tons of peaches (Prunus persica L. Batsch) were produced for shipping from Georgia during the 2002 season. The farm-gate value for the

Georgia and South Carolina peach industries combine to exceed $70 million annually.

California is the only state with a higher production level. Approximately 5670 of Georgia’s hectares are planted in bearing-age commercial peaches (NASS-USDA, 2004). Because of the warm and humid climate in the southeastern United States, it is a constant challenge to maintain tree health during and after orchard establishment. The problems that affect peach production and orchard longevity are both biotic and abiotic including fungi, bacteria, arthropods, nematodes, sunscald, frost damage, hail, drought, and even occasionally flooding. Many of the

biotic problems tend to be heightened when replanting on old peach sites (Hine, 1961; Koch,

1955; Savage et al., 1953). Good management practices (such as proper planting, fertilization,

and weed control), advances in chemical treatments (e.g., fumigants, fungicides, herbicides, and

insecticides), and increased plant vigor (resistant rootstocks and improved scion varieties) have

helped to overcome or at least reduce some of these pressures.

The two most common causes of peach tree mortality in the southeastern U.S. are Peach

Tree Short Life (PTSL) and Armillaria Root Rot (ARR). PTSL, which until recently has been

the leading cause of peach tree mortality, is the result of interactions among a number of factors, 2 including: time of pruning, improper soil pH, bacterial canker, (caused by Pseudomonas syringae pv. Syringae), and most notably the ring nematode, Mesocriconema xenoplax (Reilly et al., 1986). On the other hand, ARR is caused by fungi in the basidiomycete genus, Armillaria predominantly A. tabescens and A. mellea in the Southeast. Armillaria spp. are white-rot fungi, able to degrade the cell wall components lignin, hemicellulose, and cellulose (Peterson, 1960;

Rhoads, 1954; Savage et al., 1953). The pathogen begins by infecting bark and then progresses by moving inward to the cambium (Morrison et al., 1991). Miller (1994) found that PTSL accounted for approximately 50% of premature peach tree mortality in the southeastern U.S. while ARR caused 35% of tree mortality annually. To truly understand the importance of these diseases on the southeastern peach industry, the economic impact should be evaluated. When considering the value of lost crops based on a 15-year tree lifespan, PTSL and ARR combined accounted for a $10 million annual loss in South Carolina alone (Miller, 1994).

Okie et al. (1994) reported that PTSL losses confound those due to ARR because PTSL commonly strikes when the tree is between 3 and 6 years old, while ARR generally kills older trees. Advances in the development of nematode-resistant rootstocks (Guardian™, for example) have paved the way for replanting in PTSL sites (Beckman et al., 1998). Because of the recent developments in combating PTSL, ARR appears to be emerging as the primary soil borne pathogen of peach trees in the southeastern U.S. Because there is no commercially acceptable rootstock resistance and ARR is not limited to peach orchard replant sites, it may become a greater threat to the peach industry than PTSL.

Armillaria root rot. Armillaria (Fr:Fr) Staude, the causal organism of ARR, belongs to the phylum Basidiomycota, order Agaricales, and family Basidiomycota. Members of this genus are among the most devastating white-rot fungi in the world. Known also as the shoe-string

fungus, honey mushroom, or oak root rot fungus, Armillaria spp. attack a wide range of woody

plant species including perennials, commercial trees, forest trees, and shrubs (Hood et al., 1991;

Kile et al., 1991). Some specific plants that Armillaria spp. infect include: pine (Pinus spp.;

Ross, 1970); various conifers (Omdal et al., 1995); grapevines (Vitis vinifera; Mansilla et al.,

2001); strawberry (Fragaria spp.; Fox and Popoola, 1990); citrus (Citrus spp.), magnolia

(Magnolia spp.), camellia (Camellia japonica), walnut (Juglans spp.), sweet gum (Liquidambar styraciflua) (Raabe, 1966); cassava (Manihot esculenta; Mwenje et al., 1998); and various stone fruit crops, (e.g., Prunus persica, P. mahaleb, and P. dulcis; Raabe,1966).

In the Southeast prior to 1950, peach tree death was generally attributed to poor nutrition, winter injury, and problems associated with borers and chemical applications (Rhoads, 1954;

Savage and Cowart, 1954). With an increase in plant pathology, it was discovered that these explanations for peach tree loss were not adequate. By the mid-1950s, researchers realized that

Armillaria spp. occurred widely throughout the Southeast in both hardwood stands and peach orchards (Savage and Cowart, 1954). The species that attack peach trees are of particular concern in this region. Although A. tabescens (previously known as Clitocybe tabescens) is the primary pathogen, there is morphological evidence that A. mellea also occurs in southeastern peach orchards (Chandler and Daniell, 1982; Rhoads, 1954). For many years, these two species were regarded as one (Rhoads, 1945; Ross, 1970). However, based upon morphological characteristics such as rate of growth, rhizomorph production, and ability to luminesce, Rhoads

(1945) concluded that A. mellea and A. tabescens were indeed two distinct species.

There are three characteristic signs that are associated with infection by Armillaria. The first and most obvious characteristic is the honey-colored basidiocarps that form at the base of infected trees. The cap is generally between 5 and 15 cm wide with the underside covered by

4 loosely spaced whitish gills that radiate from the stipe (Leaphart, 1963). The spores are disseminated primarily by wind and rainfall. The two major southeastern species are easily distinguished from one another by the basidiocarp morphology. A. mellea has an annulus, a ring surrounding the stipe, while A. tabescens is exannulate. These basidiocarps may be present singularly or in clusters of more than 100 (Leaphart, 1963).

A second sign of Armillaria is the white to cream-colored fan-shaped mycelial mat. The mat extends up the tree’s phloem and cambium separating the wood from the bark, eventually leading to tree death. Small pin-point perforations are present in all stages of mycelial fan development of A. tabescens (Rhoads, 1945; Ross, 1970).

The third sign of Armillaria spp. is the melanized mycelial aggregates, called rhizomorphs, which adhere to the surface of roots, attack host tissue, and resist other soil microbes. In orchards, these ‘cords’ are predominantly associated with A. mellea. Similarly, white to brownish cords may be seen on the underside of A. mellea and A. tabescens cultures grown on Petri dishes. Presumably, these structures serve the same purpose both in vitro and in vivo, foraging for nutrients and providing a means of infection.

Another characteristic of Armillaria spp. is the ability to persist in the soil for many years. The primary means of infection appears to be the movement of actively growing roots into soil planes containing infested root pieces (Leaphart, 1963). These infected root pieces with viable mycelia or pseudosclerotia may be found in the orchard 35 years after tree removal

(Chandler and Daniell, 1982). Pseudosclerotia, which consist of swollen melanized fungal cells containing partially digested wood, form after the mycelium interacts with other fungi or host tissue. This structure shields the hyphae from soil microbes while it permits the organism to survive and complete its life-cycle when soil conditions are suitable (Fox, 2000). Studies also

suggest that Armillaria spp. have the capacity to produce antibiotic phenolics, which increase their longevity in soils (Odure et al., 1976; Vance and Garraway, 1973).

Disease symptoms are common in peach orchards and may be mistaken for a number of other biotic and abiotic problems. Above-ground symptoms of ARR include yellowing, leaf curling (along the mid-rib), leaf senescence, stunted growth, dieback of branches, and eventual

tree death. In later stages of the disease, leaves exhibit a lighter green to bronze color, limbs

located above infected roots will cease to grow and eventually die (Cooley, 1943). Generally,

trees possessing such symptoms will not survive more than 2 years. In peach orchards, groups of

dead trees tend to form in circular patterns. These patterns are indicative of the radial tree-to-tree

spread of the fungus from infection points (Chandler and Daniell, 1982). In a North Carolina

orchard trial, Chandler noted that A. tabescens did not begin to kill trees until the fourth year.

Sixty-eight percent of tree deaths caused by A. mellea occurred between the fifth and tenth year.

The pathogen has the means to live as a saprophyte on dead tissues or as a parasite on living

tissue (Leaphart, 1963). Armillaria spp. have the capability to attack healthy tissue or hosts that

have been weakened by other adverse conditions, thus making them ‘pathogens of opportunity’

(Leaphart, 1963).

The pathogenicity of Armillaria spp. varies depending on the geographic region, species,

and substrate. Researchers have tried to measure pathogenicity both in vitro and in vivo. Studies

involving inoculated potato (Solanum tuberosum L.) tubers documented the variation in

virulence and rhizomorph production of several isolates (Gregory, 1985). This study also

showed that there were marked differences among responses of young conifers and potato tubers

to the same isolates. Therefore, just because differences with response of potato tubers to

various Armillaria species were noted, this method is not a reliable test for pathogenicity studies.

Similarly, the use of cassava (Manihot esculenta) tubers for pathogenic studies was shown to be

unreliable in predicting relative pathogenicity on other host species (Mwenje et al., 1998).

While assessing Armillaria pathogenicity and virulence to seedlings of Monterey pine (Pinus

radiate), dwarf dahlia (Dahlia pinnata), and peach, Raabe (1967) noted a great degree of variability among ten A. mellea isolates. To further strengthen the case for variability among

Armillaria isolates, another study found that 11 single-spore isolates from a single basidiocarp varied widely in pathogenicity and virulence (Raabe, 1972). Although the sample size was rather small, it was suggested that these variations might help explain the large host range of

Armillaria.

Another study demonstrated that the pathogenicity of an isolate is dependent on the host genus and species (Shaw, 1977). Thomas et al. (1953) classified the varying degrees of susceptibility of a number of plant hosts to Armillaria spp. Almond (Prunus dulcis), peach, and apricot (P. armeniaca), were the least resistant while fig (Ficus carica), persimmon (Diospyros spp.), French pear (Pyrus communis), and black walnut (Juglans nigra), were the most resistant

(Thomas et al., 1953). The diversity of the specificity of Armillaria spp. varied widely. Isolates most pathogenic to hardwoods showed little if any pathogenicity toward pine seedlings. In southern England, A. mellea had the widest host range, attacking both broad-leaved shrubs as well as pines and certain conifers (Rishbeth, 1985). By contrast, A. obscura is much more specialized, killing mostly resinous conifers. Apparent pathogenicity on certain species of trees is dependent on the host response elicited by the fungus, such as gum in broadleaves or oleoresin in conifers (Rishbeth, 1985). Because of the wide range of pathogenicity within the genus,

Armillaria species are successful pathogens that are difficult to control.

There are no effective direct methods to control ARR. Once a tree shows signs and

symptoms of the disease, it is too late for treatment. Previous control methods consisted of

tactics such as proper cultural methods and girdling (Leaphart, 1963). In the mid 20th century,

the removal of diseased trees (including all root pieces with a grubbing-hoe) was an integral part of preventing disease spread (Rhoads, 1954). The fumigant carbon disulfide (also known as carbon bisulfide or CS2) may be injected into orchard soils as a means of control. While

problems associated with this treatment include incomplete control, toxicity, and corrosiveness,

it was one of the best control methods utilized in the 1950s and 1960s (Leaphart, 1963; Thomas

et al., 1953).

Soil fumigation remains a method that can give some measures of control for ARR.

Fumigation with methyl bromide is the most effective control thus far, at least on sandy soils

(Sherman and Beckman, 1999). However, its use on the heavier clay type soils typically found

in the main southeastern peach production region has generally proven unsatisfactory (Sherman

and Beckman, 1999). Regardless, the production and importation of methyl bromide will cease

in 2005 and will be completely phased out by 2015 due to its adverse environmental impact

causing ozone depletion (ARS Methyl Bromide Research website, 2004).

A second method being evaluated to lessen the impact of Armillaria spp. is the

development of resistant rootstocks. Resistant plum and peach rootstocks are being examined as

potential solutions (Beckman, 1998). Currently, plum rootstocks appear to be the most resistant

to ARR. However, peach scions on some plum rootstocks appear to suffer from problems with

graft compatibility, vigor, and productivity and will require extensive testing to assure

commercial suitability (Beckman, 1998).

One important step in developing improved management tactics is to gain a better

understanding of the diversity of the southeastern Armillaria population. Although the genus is

easily recognized and diagnosed, the species are not as easily differentiated. Understanding the

pathogenicity of southeastern isolates that infect peach may lend focus to a program for the

development of more resistant rootstock material.

Literature Review

Throughout most of the 20th century there was debate and confusion about the

classification of Armillaria spp. Until 1945, many mycologists held the common view that

Clitocybe tabescens was merely an exannulate form of A. mellea (Rhoads, 1945). However,

Rhoads (1945) concluded in his comparison between C. tabescens and A. mellea that these were

two distinctive species. At this time, Clitocybe tabescens was renamed Armillaria tabescens, It

was later determined that both A. mellea and A. tabescens exist in the southeastern U.S. (Rhoads,

1954; Ross, 1970; Savage et al., 1974).

One challenge facing researchers is to understand the diversity of the southeastern

Armillaria population. It will be of great benefit to better understand the species involved with the disease on peach as well as potential degrees of pathogenicity for various strains within these species. Throughout the years, different methods to identify and classify Armillaria have been utilized. Non-molecular approaches include isozyme and morphological analyses, as well as mating type studies. Molecular techniques such as restriction fragment length polymorphism

(RFLP) analysis appear to be more effective for differentiation at both a species and below- species levels.

Basidiocarp morphology. Several conventional techniques are employed to

discriminate among species, including examination of gross morphological characteristics. The

easiest way to distinguish between the two predominant species in southeastern peach orchards,

A. tabescens and A. mellea, is simply by the presence or absence of an annulus. While

characterization of basidiocarp morphology appears to be a straightforward approach, there are

two considerations that make this approach problematic. First, there is a very limited window of

time in which these structures appear. In the Southeast, basidiocarps are usually present for only

a few days in the fall as temperatures cool and precipitation increases. In many years,

basidiocarps will not be produced at all, even with favorable conditions. Second, separation

based on morphology alone in most cases requires a trained eye and a good deal of expertise.

Other characteristics such as ring attributes, stipe ornamentation, presence of subhymenial

clamps, basidiospore size, and basidiospore ornamentation may be helpful to differentiate

species (Watling et al., 1991).

Morphological analyses. A second means to compare species is an in vitro examination

of cultural morphology. Prior to the mating tests developed by Korhonen (1978), the three

criteria used to identify Armillaria species in culture were growth rate on 3% malt agar at 30° C, colony appearance on gallic acid agar, and the type of growth on slopes of hazelwood-malt agar

(Rishbeth, 1986). While studying English Armillaria isolates, it was revealed that the second method (growth on gallic acid) was the most reliable to distinguish species (Rishbeth, 1986).

Furthermore, these three methods were shown to be complementary and were considered concurrently to obtain valuable information. However, even a single species such as A. mellea, can vary considerably in virulence. In a similar study of in vivo characteristics, specific

differences between A. mellea and A. tabescens were found to exist with both their rhizomorph

production and their capacity to luminesce (Rhoads, 1945).

Morphological analyses are not without their problems. Not only were the characteristics of 84 pathogenic isolates (taken from individual hosts) variable, but 84 single-spore isolates taken from basidiocarps found on three different hosts were also diverse (Raabe, 1966).

Rhizomorph production and vigor of growth also differed between these two groups (cultures isolated from hosts versus single-spore isolates). A wide variation in length, number, shape, width, branching and production of rhizomorphs, colony size and shape, hyphal production where rhizomorphs break agar surface, and production of exudates were found within groups

(Raabe, 1966).

Mating experiments. Mating type studies were developed to distinguish among

Armillaria spp. in vitro. This technique is based on the knowledge that Armillaria is a group of heterothallic fungi with two (bifactorial) mating-type loci (Hintikka, 1973). In 1978, Korhonen reported that there were five sexual compatibility groups. Today, the total has expanded to nine groups known as the North American Biological Species (NABS) (Coetzee et al., 2000).

It is ideal to plate haploid unknowns with haploid tester strains so that a visible morphological transformation will occur with the exchange of genetic information. Because basidiocarps are not readily available, obtaining single-spore isolates is not always possible.

However, researchers have shown that somatic segregation of haploid isolates is possible using benomyl, a common foliar fungicide (Anderson, 1983; Proffer et al., 1987). It was hypothesized that benomyl causes abnormal segregation of whole chromosomes during mitosis (Anderson and

Yacoob, 1984). This discovery paved the way for subsequent studies to identify unknown diploid isolates.

The following is a generally accepted way of speciating unknown isolates, (Guillaumin et al., 1991):

In order to determine the species of a given isolate, an unknown isolate is plated next to a known species. If the fungi are sexually compatible, new fungal growth will exhibit diploid morphological characteristics, such as darkened crustose growth. New growth of incompatible isolates (not belonging to the same species) remains white and fluffy. This morphology, typical of haploid mycelia is expected since the incompatible fungi cannot hybridize.

In most cases, the picture is further complicated by the necessity to mate the unknown isolates with more than one tester strain because members of this genus are bifactorially heterothallic. For complete compatibility to occur, the fungi must have different alleles at each locus. If the loci contain the same alleles, they are not sexually compatible and no exchange of genetic information can occur. Therefore the growth remains characteristically haploid.

However, if one locus has matching alleles and the other has different alleles, there are varying degrees of semi-compatibility that may occur which further complicate interpretation of the results. Methods of cultural identification are time consuming, require a great deal of expertise, and are not particularly reliable (Guillaumin et al., 1991; Perez-Sierra et al., 2000).

Mating experiments are also convoluted by the possibility that hybridization may actually occur among A. mellea and A. tabescens isolates in the Southeast. Based on molecular analysis of the rDNA region, Bryson et al. (2003) have shown that certain isolates exhibit banding patterns that are characteristic of both Armillaria spp. While this does not make the mating studies invalid, it does add another dimension that must be considered.

Isozyme analysis. Another method used to classify Armillaria isolates is isozyme analysis. Isozymes, different molecular forms of an enzyme, have the same enzymatic function but possess slightly different net charges, configurations, or sizes. Because of these differences,

the molecules migrate at different rates during gel electrophoresis which allows them to be

separated and visualized. The resulting banding patterns act as “fingerprints” for species as well

as isolates. Isozyme analysis is both an easy and inexpensive way to identify Armillaria species

(Bergmann et al., 1996; Perez-Sierra et al., 2000; Robene-Soustrade and Lung-Escarmant,

1997). However, there are problems with this technique. Two separate studies found that the

rate of polymorphisms was rather low. In one study, only three of 20 enzymes assessed in

Armillaria spp. exhibited polymorphisms (Lin et al., 1989). Guillaumin et al. (1996) found that although isozyme analysis was reliable in characterizing species, it was not as useful in determining polymorphisms among different isolates of the same species. Most of the seven enzymes examined showed clear differences between the two species, A. gallica and A. cepistipes. However, Guillaumin et al. (1996) noted that this method was still limited by the low number of polymorphisms.

Molecular techniques. Currently, the most prevalent and reliable method of differentiating among genera, species, and even races of an organism is by examining its genotype. With the advent of the polymerase chain reaction (PCR) in 1983, many different techniques for studying an organism’s genome with relative ease have been developed. Kary

Mullis and his co-workers developed this new technology as a way to amplify specific segments of DNA (Edel, 1998). PCR allows for specific and sensitive detection and production of DNA fragments from numerous sources (Foster et al., 1993).

DNA extraction. The first important step in molecular characterization is the acquisition of an adequate amount of intact, pure, or high-quality, DNA from fungal cells. Previous studies have revealed a number of techniques. They involve a range of methods for tissue disruption as well as a variety of chemical constituents involved in the actual procedure. A common method

13 of extraction starts with 50 mg of mycelia (Raeder and Broda, 1985). The fungal DNA is first extracted in a buffer containing sodium dodecyl sulfate (SDS), ribonuclease A (RNAse A), phenol, and NaCl. After chloroform extraction and another RNAse A treatment, the DNA is precipitated with isopropanol. DNA was isolated easily without the presence of contaminants that may interfere with PCR. Raeder and Broda (1985) found that the entire procedure took approximately 1.5 to 2 hr and yielded DNA that was 0.1 to 0.15% of the initial quantity.

The characteristic polyphenol production in cultures of southeastern Armillaria isolates appears to decrease DNA yield and quality. Polyphenol oxidases cause the oxidation and polymerization of compounds that are potentially toxic to the fungus (Garraway et al., 1991).

These compounds are evident in the marginal browning of oxidized tissues as well as the browning of media in vitro. Polyphenolic oxidases inactivate enzymes, precipitate proteins, and form cross-links with nucleic acids, which in turn inhibit most molecular manipulations (Koonjul et al., 1998). However, DNA extraction buffer can be amended with polyvinylpyrrolidone

(PVP) to bond with and inhibit the polyphenols (Chirgwin et al., 1979). Using PVP in DNA extractions of cotton (Gossypium hirsutum), known for large quantities of polyphenol production, 0.5mg DNA per g of tissue was collected (John, 1992). A midiprep method for isolating DNA from plants containing high phenolic levels such as field bean (Vicia faba), tomato (Lycoperscion esculentum), and potato was developed by Pich and Schubert (1993).

Using the midiprep method, 0.02 to 0.04% of high-quality DNA can be obtained from the original leaf and stem tissue (Pich and Schubert, 1993).

In developing their PCR identification technique for Armillaria spp., Harrington and

Wingfield (1995) used two different approaches to acquire template DNA. They not only used the Lee and Taylor (1990) extraction method, which yielded high-quality DNA from filamentous

fungi, but they also utilized a novel time-saving technique referred to as direct amplification. A

pipette tip was scraped approximately 1 cm across the periphery of the colony and then dipped

into a tube containing the PCR reaction mixture. The suspended mycelia were placed directly in

the PCR reaction for amplification.

RAPD analysis. Random amplified polymorphic DNA (RAPD) analysis, also called arbitrarily primed PCR, is a method based on the selective amplification of genomic sequences that, by chance, are flanked by adequate matches to an arbitrarily chosen primer (Welsh et al.,

1995). RAPD primers are shown to be effective tools that differentiate among fungi at a generic

(Tommerup et al., 1995), specific (Shi et al., 1995), and even sub-specific level (Crowhurst et al., 1991; Tommerup et al., 1995; Voight et al., 1998). These techniques have been employed to determine the efficacy of fungi as biological control agents (Schena et al., 1999) and to understand the varying degrees of resistance to fungicides (Delye et al., 1997).

Few investigations have characterized Armillaria spp. by RAPD analysis. While comparing the genotypes of 73 Armillaria isolates (A. ostoyae, A. gallica, and A. cepistipes), one

study revealed that RAPD analysis showed greater genetic variations than other molecular

techniques (Guillaumin et al., 1996). A study on the variability with A. ostoyae found that

RAPD analysis was a valuable tool (Schulze et al., 1997). Advantages of RAPD analysis include: 1) the brief amount of time required for completion, 2) the capacity to perform this analysis without having prior knowledge of the genomic sequence, and 3) the ability to provide markers in regions not accessible with random fragment length polymorphism (RFLP) analysis

(Williams et al., 1990).

Ribosomal DNA. Ribosomal DNA is commonly used to understand the relationships among many genera of fungi. Ribosomes are cytoplasmic bodies composed of protein and

ribosomal RNA. They are the sites of nonspecific protein synthesis (Gardner, 1975). In most eukaryotic nuclei, the rRNA genes are in tandem clusters, also called rDNA (Lewin, 1990).

Because the rDNA, found universally in living cells, has such an important function, its evolution may be used to study the development of the whole genome (Edel, 1998). This region of DNA contains both variable and conserved areas that may be useful in making comparisons and discriminating among genera, species, and subspecies (Edel, 1998). The nuclear rDNA of fungi is one of the most highly conserved regions in the genome (Hibbett, 1992).

Most fungi contain a single transcription unit of nuclear rDNA organized from 5’ to 3’ with one 18S small sub-unit (SSU), followed by one 5.8S gene, and one 25-28S large sub-unit

(LSU) (Figure 1). This three-gene sequence may be repeated tandemly multiple times.

Depending on the fungal taxa being examined, another gene, the 5S unit, may or may not be included in the repeated region. Two internal transcribed spacers (ITS-1 and ITS-2) separate the genes within a single three-gene unit and the intergenic spacer (IGS) separates each set of rDNA genes into repeated units (Figure 1).

Because the 18S gene evolves slowly, it is useful in making comparisons among distantly related organisms. However, the IGS and ITS regions appear to have evolved much quicker, making them much more variable. Therefore, these areas are useful in comparing fungal species within a genus or even strains within species (Anderson and Stasovski, 1992; Baumgartner and

Rizzo, 2001; Edel, 1998; Harrington and Wingfield, 1995; Perez-Sierra et al., 1999).

ITS amplification. The fungal ITS region of rDNA contains less genetic variability than the IGS region in the same operon (Lee and Taylor, 1992; Schulze et al., 1997). Using the primer

ITS-1 (White et al., 1990), the ITS region was found to be nearly uniform among the 24 strains of Armillaria examined by Anderson and Stasovski (1992). Although these authors discovered

that there was more variation between A. mellea and A. tabescens, they concluded that the ITS

was too uniform to characterize the majority of the isolates studied and too variable to resolve the relationships among other species (Anderson and Stasovski, 1992). However, other researchers have concluded that the ITS region is useful for identification at the species level

(Lee and Taylor, 1992; Schulze et al., 1997).

While studying 20 Armillaria isolates from Picea spp., Alu I digestion was shown to inadequately define the ITS region at an intraspecific level (Schultze et al., 1997). Only through their further work with RAPD primers did they discover that there were indeed intraspecific differences in the genomic DNA (Schultze et al., 1997).

IGS amplification. One of the most common techniques to characterize DNA sequences is restriction analysis. After a short region of DNA is amplified using PCR, the restriction endonuclease (type II) is used to cleave the PCR amplicon at a specific sequence (usually between 4 and 8 bp) (Perez-Sierra et al., 2000). The fragments are then visualized using gel electrophoresis to quantify their size.

Harrington and Wingfield (1995) found that by amplifying the IGS region of various

Armillaria spp. and then restricting with endonucleases they were able to differentiate among species. To amplify the IGS-1 region they used the primers LR12R,

5’CTGAACGCCTCTAAGTCAGAA3’ (Veldman et al., 1981) and O-1,

5’AGTCCTATGGCCGTGGAT3’ (Duchesne and Anderson, 1990). With only three distinct band sizes representative of the eleven species examined, amplification alone was not informative. However, digestion with Alu I proved to distinguish among species more effectively (Harrington and Wingfield, 1995). The three species that were not clearly defined with Alu I (A. ostoyae, A. gemina, and A. borealis) were distinguished successfully with Bsm I,

Nde I, and Hind II. In addition, these techniques were used to characterize Armillaria spp. in

particular geographic regions (Baumgartner and Rizzo, 2001; Mwenje et al., 2003),

Similarly, these methods of amplification and restriction were used to distinguish among

European isolates of Armillaria (Perez-Sierra et al., 1999). A. mellea, A. tabescens, and A. gallica were clearly defined using the enzyme Alu I. A. ostoyae, A. borealis, and A. cepestipes were differentiated using Bsm I. Additionally, previously unreported patterns for A. mellea and

A. gallica were discovered. By using the techniques of Harrington and Wingfield (1995), five species (A. ostoyae, A. sinapina, A. nabsnona, A. gallica, and NABS XI) that affected seven different hosts were identified on the Olympic Peninsula of Washington state (Banik et al.,

1996).

One study attempted to use nucleotide sequences of the ITS and IGS regions as a means of speciation (Sicoli et al., 2003). Primers designed for A. mellea proved to be species-specific for both the ITS and IGS regions. A. tabescens and A. mellea were both divergent enough to

exhibit differences in the IGS region. However, the similarities between A. borealis and

A.ostoyae and between A. cepistipes and A. gallica were such that these rDNA regions were not

useful for speciation (Sicoli et al., 2003).

A number of unknown isolates in Pennsylvania were identified by comparing their

restriction patterns with the patterns of six known Armillaria species (Frontz et al., 1998). By

using several different restriction enzymes (Hpa I, Hae III, Alu I, and Hinf I), 58 unknown

isolates found in the Tioga State Forest were classified at the species level.

Examination of isolates of the newly characterized A. nabsnona showed three different

restriction patterns when the IGS region was cleaved with Alu I (Volk et al., 1996). These

results demonstrate that inferences about relationships among isolates at a level below species

18 can also be made using this technology. When examining the relatedness between four North

American species (A. cepestipes, A. sinapina, NABS X, and NABS XI) with Alu I, it was shown that not only did a single species exhibit multiple banding patterns, but multiple species shared a common banding pattern. From these and other studies, it is apparent that restrictions of the rDNA may be useful in further understanding the diversity within a given species (Chillali et al.,

1997; Kim et al., 2000; Schulze et al., 1995; White et al., 1998). Despite the success of this method, there are some disadvantages. Restriction analysis adds an additional day to the time required to complete the procedure and requires large amounts of PCR product (Foster et al.,

1993; Perez-Sierra et al., 2000).

The primary objective of this research is to gain a better understanding of the inter- and intraspecies variability of Armillaria spp. that infect southeastern peach orchards. By using molecular techniques with RAPD primers, specific primers previously identified for Armillaria spp., and restriction analysis of the ITS and IGS regions of the rDNA, a better understanding of the unknown southeastern isolates may be achieved.

Figure 1: Schematic illustration of the typical arrangement of rDNA of basidiomycetes, including primer locations. The internal 19 transcribed spacer (ITS) and the intergenic spacer (IGS) regions are shown with primer annealing sites (not to scale).

CHAPTER 2

MATERIALS AND METHODS

Isolate collection. During the fall of 2000, research and extension personnel from

Georgia and South Carolina joined in a collaborative effort to establish a representative regional

collection of isolates of Armillaria spp. In the initial collection, 52 isolates were obtained from

peach orchards throughout South Carolina. Basidiocarps and mycelial mats from the crown and

upper roots of the tree were gathered. In the course of the next three years, additional isolates

were obtained from peach orchards in Georgia and Alabama. Currently, there are 77 isolates

included in the active collection.

Because there was some duplication at several of the collection sites, only a subset of the

collection was examined in this study. In some cases, more than 15 isolates were collected at a

single location. In other instances, three samples were taken from the same tree (basidiocarp,

mycelial mat from trunk, and root piece). To reduce the number of isolates to a more

manageable size and to decrease the duplication of a certain isolate or a particular location, a group of 39 isolates was examined in this study (Table 1, Fig. 2). There were no single-spore, haploid isolates represented among the unknowns.

In addition, known isolates ST5 (A. mellea) and TA1 (A. tabescens), acquired from Dr.

Guido Schnabel at Clemson University, were also analyzed as a means of comparison. ST5 was isolated in Virginia. The species was identified by the USDA Forest Center for Forest Mycology in Madison Wisconsin (Kim et al., 2000). TA1 was isolated from Quercus nigra in Florida 21 during 1988. It was identified as a haploid tester strain by the Schnabel lab (personal communication, 2002).

Isolation and maintenance. After the samples had been collected they were transported on ice to the USDA-ARS Southeastern Fruit and Tree Nut Research Laboratory in Byron, GA, where they were stored at 4 to 6ºC until isolation. After excess soil was removed by washing with tap water, plant material and basidiocarps were surface-disinfected with 95% ethanol. The outside of woody tissue was flamed to remove any potential contaminants. Both the basidiocarps and woody tissue were broken open to expose Armillaria mycelia. Very small portions of the fungal tissue were teased away from plant tissue using forceps and plated on

MYEA (2% malt extract, 0.2% yeast extract, 1.5% agar) plates amended with streptomycin (100

µg/mL) with rescue from contaminants until isolated as a single culture. For routine maintenance and storage, isolates were sub-cultured every 3 to 4 wk and maintained at 28ºC on

MYEA plates in the dark. For long-term storage, cultures were grown on MYEA slants modified with autoclaved sawtooth oak (Quercus acutissima) acorns as a carbohydrate source and stored at 4ºC.

DNA extraction. Because of initial difficulties, including poor repeatability of extractions and problems associated with polyphenolic accumulation, two different DNA extraction techniques were explored. One method, described by Harrington and Wingfield

(1995), involves loading mycelia directly into the PCR amplification mixture. Under a sterile laminar flow hood, a pipet tip was scraped ca. 1 cm across the periphery of an actively growing colony. The tip was then swirled in a 0.2 mL tube containing the pre-mixed PCR reagents to dislodge the mycelia. Presumably, the heat of the denaturation step of PCR (95º C for 2 min) was sufficient to release the DNA by lysing the fungal cells. Mycelia of a quantity visible to the

22 eye, were considered too much for successful amplification (G. Schnabel, personal communication, 2001).

The second extraction method combined a rapid extraction protocol with the addition of polyvinylpyrrolidone (PVP) to bind phenolics (John, 1992; Lee and Taylor, 1990; Pich and

Schubert, 1993). Isolates were grown in both MYE liquid culture and on MYEA plates. For the liquid culture, 250 mL Erlenmeyer flasks containing 50 mL MYE were inoculated with 4 to 8 agar plugs (ca. 5x5mm) from actively growing plates. These cultures were shaken in the dark at

120 rpm for 2 to 3 wk at room temperature (23-25ºC) on a benchtop shaker. The contents of the

250 mL flasks were then poured into a Waring blender and macerated at approximately 400 rpm for 10 to 20 sec at room temperature. The chopped tissue and media were poured aseptically into a 500 mL flask containing 100 mL MYE. The mixture was then shaken (Innova 4300 incubator shaker, New Brunswick Scientific, Edison, NJ) at 120 rpm at 28ºC for 1 to 2 months. The cultures were strained with 2-ply cheesecloth and blotted with paper towels to remove excess moisture. The mycelia were then transferred to 25 mL Falcon tubes and freeze-dried overnight in a centrifugal evaporator (RC 1022, Jouan, Winchester, VA). Finally, each isolate was frozen in liquid nitrogen and macerated in a mortar with a pestle before extended storage at -20ºC. For

DNA extraction, ca. 300 mg of thawed mycelial tissue was placed in a 2 mL microfuge tube with

900 mL extraction buffer (50 mM Tris-HCl pH 7.2, 50 mM EDTA, 3% SDS).

Alternatively, fresh mycelia grown on MYEA petri plates (100 x 15mm) were used for

DNA extraction. The cultures were maintained in incubation ovens (26 to 28ºC) for 3 to 4 wks.

Using a scalpel, mycelia were dissected away from the periphery of the culture and ca. 300 mg weighed. The mycelia were placed in a 2 mL microfuge tube with 900 µL extraction buffer and ground for 20 sec with a pellet pestle (Kontes, Vineland, NJ).

Whether liquid or solid media was used, the remainder of the extraction process was the same. To complete the extraction mixture, β-mercaptoethanol and PVP (stock solution: 32%

MW 40,000 polyvinylpyrrolidone, stored at 4ºC) were added to the tissue to achieve a final concentration of 1.0 and 6.0%, respectively, and vortexed for 30 sec. The mixture was then placed in a 68ºC water bath for 30 min. Next, 0.1 volume of 5M potassium acetate was added and gently inverted to mix. The tube was then transferred to an ice bath for 30 min. To precipitate the cell debris, the suspension was centrifuged at 4ºC for 10 min at 16,000 x g

(Eppendorf 5415 C, Westbury, NY). After the supernatant was transferred to a fresh tube, 0.6 volumes of isopropanol was added, gently mixed and centrifuged at 4ºC (16,000 x g) for 15 min to precipitate DNA. The supernatant was then poured off and the tube was inverted on a paper towel to dry.

Following resupension of the pellet in 200 µL TE buffer (10 mM Tris Cl, pH 7.4; 1 mM

EDTA, pH 8.0), 0.33 volume of 5M sodium chloride was added and mixed gently. After centrifugation (16,000 x g) at 4ºC for 20 min, the supernatant was transferred to a fresh tube and

0.6 volumes of 100% isopropanol were added. To aid with DNA precipitation, the solution was held at -20ºC for 5 to 10 min. After gently mixing, the tube was centrifuged (16,000 x g) at 4ºC for 15 min. The supernatant was poured off and tube inverted to air dry. The pellet was washed twice with 100 µL of 70% EtOH and centrifuged at 4ºC (16,000 x g) for 10 min. The supernatant was poured off and the tube again inverted to air dry. The final drying step was performed under vacuum for 30 to 45 min to remove excess EtOH. Once completely dehydrated, 100 µL TE buffer was added to the pellet for resuspension overnight at 4ºC. The resuspension was completed the following day with gentle inversion several times. Finally, the

DNA content was quantified spectrophotometrically at 260 nm (Biorad SmartSpec 3000,

Hercules, CA).

Whole-genome analysis. RAPD analysis was used to determine whether the unknown isolates exhibited any distinguishing patterns. This experiment was approached in two different ways. Initially, UBC (Biotechnology Laboratory, University of British Columbia, Vancouver,

BC, Canada) primer set #5 (401-500; Table 2) was run with isolate csf1. Additionally, the two

‘known’ isolates, ST5 (A. mellea) and TA1 (A. tabescens), along with nine unknown isolates

(usd 2, ync 3, ctn 1, pch 3, blk 2, gmg 1, gmg 2, csf 2, dky 1) were analyzed using UBC primers

401-500.

All RAPD procedures contained the same PCR reagents and were run under similar cycling conditions. The following components were included in the PCR reactions: 5 ng template DNA, PCR buffer at a final concentration of 1x (supplied with enzyme), 4 mM MgCl2,

250 µM dNTPs, 1 unit Taq, and 0.2 µM RAPD primer per 25 µl reaction. Using a GeneAmp

PCR System 9700 (Perkin Elmer Applied Biosystems, Foster City, CA) the reaction was denatured for 2 min at 94ºC, followed by 35 cycles of 1 min at 94ºC (denaturation), 1 min at

36ºC (annealing), and 2 min at 72ºC (elongation). The final elongation step was 72ºC for 7 min followed by a 4ºC hold.

IGS analysis. Each unknown isolate, along with ST5 and TA1, was amplified with

LR12R and O-1 primers (Table 2) as described previously (Baumgartner and Rizzo, 2001;

Harrington and Wingfield, 1995; Mwenje et al., 2003). Following amplification, the bands were restricted with Alu I (Harrington and Wingfield, 1995). The PCR product (6 µL) was combined with 6 units of Alu I and 0.6 µL of 10x buffer (provided with enzyme). The digestion proceeded at 37ºC for 1 to 3 hr, prior to agarose gel electrophoresis.

Primer sets that were previously designed to distinguish among European isolates of A. tabescens and A. mellea were also included in the study. ATA-1 and ATA-2 were developed to detect A. tabescens, while AME-1 and AME-2 were developed to identify A. mellea (Table 2)

(Sicoli et al., 2003).

Both IGS analysis methods, those developed by Harrington and Wingfield and Sicoli et al., were carried out under the same conditions, including concentration of PCR reagents and

PCR cycling parameters. A 25 µl reaction contained: 5 ng template DNA, 10x PCR buffer diluted to 1x for the reaction (supplied with enzyme), 4 mM MgCl2, 250 µM dNTPs, and 1 unit

Taq polymerase. The primer concentration was 12.5 mM per reaction. The initial denaturation step was 95ºC for 2 min, followed by 35 cycles of 30 sec at 90ºC (denaturation), 40 sec at 60ºC

(annealing), and 30 sec at 72ºC (elongation). The final elongation step was 72ºC for 7 min followed by a 4ºC hold.

ITS analysis. Because the ITS region is less heterogeneous than the IGS, it was also considered as a means of species characterization. Bryson et al. (2003) developed primers based on sequences of several Armillaria isolates found throughout the United States. AmITS-1,

AtITS-1 and the universal primer ITS-2 were used to differentiate between southeastern isolates of A. tabescens and A. mellea (Table 2). The protocol is the same as that used in the examination

of the IGS region, with the exception that three (two forward and one reverse) primers were being used for each PCR instead of two.

Additionally, the universal primer set ITS-1 and ITS-4 (Table 2) was used to amplify

through the ITS-1, 5.8S, and ITS-2 regions of the rDNA. The PCR reagents and cycling

parameters were the same as in the amplification of the IGS (mentioned above). Because the

ITS is a highly conserved region, nucleotide sequences were chosen from the National Center for

Biotechnology Information (NCBI) database (2004) and analyzed for potential restriction sites with the Nebcutter web server (Nebcutter v. 2.0, http://tools.neb.com/NEBcutter, 2004). Isolates

U54818 (A. mellea) and U54821 (A. tabescens), submitted to the database by Chillali et al.

(1998), were selected for analysis. Mbo II was chosen because of its apparent potential to cleave

U54821 at two positions, 268 and 608 bp, while the ITS of U54818 was restricted only once at

280 bp. Similarly, according to Nebcutter (2004) analysis, Hha I should restrict at one site for

U54821 (330 bp) while U54818 should be restricted in two positions (346 and 654 bp).

After amplification, the products were digested with Mbo II and Hha I to determine if polymorphisms within a given species were present in this region. The Mbo II and Hha I digestions were carried out with the same reagents (5 µl of ITS PCR reaction, 0.2 µl restriction enzyme, and 1.25 µl buffer provided with enzyme) with the exception of the addition of 0.5 µl 1x

BSA in the Hha I restrictions. These digestions were carried out for 1 hr at 37ºC.

Visualization of banding patterns. Amplified PCR products (6 µL combined with 1 µL gel-loading buffer: 0.25 % bromophenol blue, 0.25 % xylene cyanol FF, 30 % glycerol) were visualized on a 2 % agarose gel (Ultrapure Agarose 1000, Invitrogen, Carlsbad, CA). Gels were made with 0.5x TBE buffer (5x stock solution: 54 g Tris base, 27.5 g boric acid, 20 mL EDTA

(0.5 M, pH 8.0) in 1L). Prior to casting, 1.5 mg of ethidium bromide was added to the gel. As a standard measure of DNA fragment sizes, 100 bp DNA Ladder (Promega, Madison, WI) was used.

Depending on the number of reactions being examined, either a small gel (8.5 x 6 x 0.5 cm) or a large gel (25 x 20 x 0.5 cm) was used. Both gels were run at 3 milliamps (223 V for the large, 125 V for the small) for 1 hr. Finally, the bands were visualized at 302 nm. The images were captured digitally with an Olympus MagnaFire SP camera (model S99810, Olympus

America, Melville, NY) equipped with a 7000 Navitar TV zoom lens. The DNA band sizes were quantified with Gel-Pro Analyzer Software (Media Cybernetics, Silver Spring, MD).

Table 1: Armillaria isolates collected from southeastern peach orchards and used in the current study.

NAME COUNTY STATE REGION SOURCE

ben 1 York South Carolina Piedmont Trunk ben 2 York South Carolina Piedmont Trunk ben 3 York South Carolina Piedmont Basidiocarp blk 1 York South Carolina Piedmont Trunk blk 2 York South Carolina Piedmont Trunk cly 1 Spartanburg South Carolina Piedmont Basidiocarp cly 2 Spartanburg South Carolina Piedmont Root cly 3 Spartanburg South Carolina Piedmont Trunk cly 4 Spartanburg South Carolina Piedmont Basidiocarp cly 5 Spartanburg South Carolina Piedmont Trunk cly 6 Spartanburg South Carolina Piedmont Basidiocarp cly 7 Spartanburg South Carolina Piedmont Trunk csf 1 Chesterfield South Carolina Ridge Trunk csf 2 Chesterfield South Carolina Ridge Basidiocarp csf 3 Chesterfield South Carolina Ridge Trunk ctn 1 Chilton Alabama Ridge Trunk dky 1 Crawford Georgia Piedmont Trunk gmg 1 Spartanburg South Carolina Piedmont Trunk gmg 2 Spartanburg South Carolina Piedmont Trunk hlm 1 Edgefield South Carolina Ridge Basidiocarp hlm 2 Edgefield South Carolina Ridge Root hls 1 Saluda South Carolina Ridge Root jns 1 Edgefield South Carolina Ridge Basidiocarp mlr 1 York South Carolina Piedmont Trunk mlr 2 York South Carolina Piedmont Trunk pch 1 York South Carolina Piedmont Trunk pch 2 York South Carolina Piedmont Trunk pch 3 York South Carolina Piedmont Trunk stc 1 Edgefield South Carolina Ridge Basidiocarp swl 1 Chilton Alabama Ridge Trunk tms 1 Oconee Georgia Piedmont Root tms 2 Oconee Georgia Piedmont Trunk tms 3 Oconee Georgia Piedmont Trunk ttn 1 Saluda South Carolina Ridge Root usd 1 Peach Georgia Piedmont Trunk usd 2 Peach Georgia Piedmont Trunk ync 1 Edgefield South Carolina Ridge Trunk ync 2 Edgefield South Carolina Ridge Trunk ync 3 Edgefield South Carolina Ridge Trunk

Table 2: Polymerase chain reaction (PCR) primers used to amplify regions of Armillaria DNA.

PRIMER SEQUENCE REFERENCE Ame-1b AAG AAT CAT GAG ATA TCA TCA GT Sicoli et al., 2003 Ame-2b TTA GAA AAT CCG CCT TAG AAA C Sicoli et al., 2003 AmITS1a CTG TTG CTG ACC TGT TA Bryson et al., 2003 Ata-1b TTG CCT TGA ACC CTG TTA TAA GGC Sicoli et al., 2003 Ata-2b TGC CAA AAT CGT TGC ACG CCG C Sicoli et al., 2003 AtITS1a GAA GGG TTG CTT TCG AGC TC Schnabel et al, in review ITS1a TCC GTA GGT GAA CCT GCG G www.biology.duke.edu/fungi/ ITS2a GCT GCG TTC TTC ATC GAT GC Bryson et al., 2003 ITS4a TCC TCC GCT TAT TGA TAT GC www.biology.duke.edu/fungi/ LR12Rb CTG AAC GCC TCT AAG TCA GAA Harrington and Wingfield, 1995 O-1b AGT CCT ATG GCC GTG GAT Harrington and Wingfield, 1995 UBC #5c #’s 401-500 Univ. of British Columbia a Amplifies ITS region of rDNA b Amplifies IGS region of rDNA c Random primer, amplifies across entire genome

Code County Isolates AL-1 Chilton ctn, swl GA-1 Oconee tms GA-2 Crawford dky GA-3 Peach usd SC-1 Spartanburg cly, gmg SC-2 York ben, blk, mlr, pch SC-3 Chesterfield csf SC-4 Saluda hls, ttn SC-5 Edgefield hlm, jns, stc, ync

Figure 2: Geographical distribution of Armillaria isolates collected from southeastern peach orchards. 30

CHAPTER 3

RESULTS

DNA extraction. Initially, while protocols were being finalized for this research, a representative group of eight isolates was examined. Using the direct amplification method

(Harrington and Wingfield, 1995), two of the eight isolates did not produce any bands after repeated attempts. Given an overall success rate for direct amplification below 20%, it was concluded that the method was not suitable for thorough analysis of the southeastern set of

Armillaria spp. isolates.

The second extraction method, involving the protocol of Lee and Taylor (1990) combined with the inclusion of PVP (John, 1992; Pich and Schubert, 1993), proved to be a more consistent way to extract intact, pure genomic DNA. Although liquid culture was preferred if large quantities of tissue were required, younger, actively growing mycelia on the periphery of cultures grown on MYEA plates was the most reliable for DNA extraction. The PVP extraction of actively growing cultures from agar using the Lee and Taylor (1990) method routinely yielded high-quality DNA between 0.01 and 0.02% of the initial material, and this DNA performed efficiently as PCR template.

Whole-genome analysis. Attempts were made to amplify isolate csf1 with UBC primer set #5 (primers 401-500). Of these reactions, approximately half (53 out of 100) of the primers produced a banding pattern. Most of the patterns were too complex and nondiscriminant to indicate which primers might be good candidates for isolate classification. Isolates ST5 (A. mellea), TA1 (A. tabescens), and nine unknown isolates (usd 2, ync 3, ctn 1, pch 3, blk 2, gmg 1,

32 gmg 2, csf 2, dky 1) were then analyzed using the 53 UBC primers from set #5 that did produce banding patterns. While many patterns were produced using the RAPD approach, no patterns were noted that distinguished species (data not shown).

IGS analysis. In order to classify the isolates based on previously described banding patterns (Harrington and Wingfield, 1995), the IGS region was PCR-amplified and restricted with Alu I. Upon amplification with LR12R and O-1, all 39 isolates, as well as TA1 and ST5 produced a band of approximately 825 bp (Figs. 3, and 4). Following restriction with Alu I, each unknown isolate fell into one of three groups (Table 3, Fig. 4). The first group, representing the majority of the isolates, exhibited a double band at approximately 470 and 430 bp, as well as a single band at 240 bp. This group included five of the Georgia isolates (tms 1, tms 2, tms 3, usd

1, and usd 2), one of the Alabama isolates (swl 1), 17 of the South Carolina isolates (ben 1, ben

2, ben 3, blk 1, blk 2, cly 7, hlm 1, hlm 2, hls 1, jns 1, mlr 1, mlr 2, pch 1, pch 2, pch 3, stc 1, and ttn 1), and the known A. tabescens isolate TA1. Another banding pattern found among these isolates consisted of the same bands mentioned above (470, 430, and 240 bp) with the addition of an intermediate band of approximately 325 bp. This group was composed of one Alabama isolate (ctn 1), five South Carolina isolates (gmg 1, gmg 2, ync 1, ync 2, and ync 3), and the

Georgia isolate dky 1. The third pattern exhibited a doublet of sizes 535 and 480 bp and a single band at approximately 180 bp. This group included nine unknown isolates from South Carolina

(csf 1, csf 2, csf 3, cly 1, cly 2, cly 3, cly 4, cly 5, and cly 6) and the known A. mellea isolate

ST5.

Another approach was needed to assess whether the intermediate group was A. tabescens or A. mellea or possibly a hybrid of the two species as suggested by Bryson et al. (2003). A subsequent approach sought to identify the isolates by using primers developed from the IGS

33 region of A. mellea and A. tabescens from Europe (Sicoli et al., 2003). Upon amplification with primers Ata-1 and Ata-2, 30 of the isolates, as well as TA1, displayed either a single or double band (Figs. 3, and 5). Isolates ben 1, ben 2, blk1, cly7, ctn 1, dky 1, gmg 1, gmg 2, hlm 2, mlr 2, pch 3, stc 1, swl 1, tms 2, tms 3, ttn 1, usd 1, ync 1, and ync 2 exhibited a doublet pattern with bands at approximately 390 and 360 bp. Isolates ben 3, blk 2, hlm 1, hls 1, jns 1, mlr 1, pch 1, pch 2, tms 1, usd 2, ync 3, and known A. tabescens isolate TA1 exhibited a single band at approximately 385 bp. There was no amplification in any of the isolates with the primers Ame-1 and Ame-2, which were developed for the IGS region of the European A. mellea isolates (Table

3, Figs 3, and 5). Isolates csf 1, csf 2, csf 3, cly 1, cly 2, cly 3, cly 4, cly 5, cly 6, and known A. mellea isolate ST5 did not amplify with either Ata or Ame primer set.

ITS analysis. By using primers AtITS1, AmITS-1, and ITS-2, further affirmation of species was sought. Thirty of the 39 isolates exhibited a single band at approximately 150 bp.

Isolates representative of ben 1, ben 2, ben 3, blk 1, blk 2, cly 7, hlm 1, hlm 2, hls 1, gmg 1, gmg

2, jns 1, mlr 1, mlr 2, pch 1, pch 2, pch 3, stc 1, ttn 1, ync 1, ync 2, and ync 3 (South Carolina); ctn 1 and swl 1 (Alabama); dky 1, tms 1, tms 2, tms 3, and usd 1, usd 2 (Georgia); as well as

TA1 were included in this group (Table 3, Figs. 3, and 6). Seven of the remaining nine isolates produced a doublet banding pattern of approximately 300 and 270 bp. This group included csf

2, csf 3, cly 2, cly 3, cly 4, cly5, and cly 6 of South Carolina. Isolates csf 1, cly 1, and ST5 produced a single band at approximately 270 bp.

A final approach to understanding the southeastern Armillaria isolate set involved the amplification with the universal primers ITS-1 and ITS-4 and restriction analysis. All isolates produced a band of approximately 850 bp (Figs. 3, and 6). After restricting the product with

Mbo II and Hha I, each enzyme yielded several distinct patterns among the isolates (Table 3).

When restricted with Hha I, the ync 1, ync 2, and ync 3 isolates produced a pattern of 520 and

215 bp (Fig. 7). Twenty-seven unknown isolates as well as the known A. tabescens isolate TA1 produced bands of 470 and 390 bp when restricted with Hha I (Figs. 3, and 7). This group included cly 7 and all isolates from other sites with the exception of ync, cly, and csf. Csf 1 produced bands of 500, 475, 400, and 295 bp when restricted with Hha I (Fig. 7). Isolates csf 2, csf 3, cly 3, and cly 4 each exhibited restriction patterns of 455, 400, and 295 bp (Fig. 8).

Isolates cly 3, cly 5, and cly 6 revealed bands of 510, 460, 410, and 320 bp. Isolate cly 1 had the simplest restriction pattern with bands at 410 and 295 bp. The known A. mellea isolate ST5 was not restricted with Hha I.

Isolates ync 1, ync 2, and ync 3 produced a pattern with bands at 870, 530, and 340 bp when restricted with Mbo II (Fig. 7). TA1 and 27 unknown isolates produced a pattern with bands at 870, 530, 355, and 340 bp (Figs. 3, and 7). This group included cly 7 and all isolates from other sites with the exception of ync, cly, and csf. Mbo II was not a useful enzyme for classifying isolates csf 1, cly 1, csf 2, csf 3, cly 2, cly 3, cly 4, cly 5, or cly 6. Because it restricted the DNA at sites too numerous to allow discrimination of species Mbo II restrictions data were not included in these results. Similarly, Mbo II did not produce a successful restriction of the known A. mellea isolate ST5.

Table 3: Summary of the molecular analysis of known and unknown Armillaria isolates.

ISOLATEa LR12R LR12R ATA-1 AME-1 AtITS1 ITS1 ITS1/ ITS1/ O-1 O-1 ATA-2 AME-2 AmITS1 ITS4 ITS4 ITS4 (bp)b Alu I (bp)d (bp)e ITS2 (bp)g Hha I Mbo II (bp)c (bp)f (bp)h (bp)i ST5 825 535,480,180 -- -- 270 850 ^ ^ TA1 825 430,240 385 -- 150 850 470,390 870, 530, 355, 340 csf 1 825 535,480,180 -- -- 270 850 500,475,400,295 ^ cly 1 825 535,480,180 -- -- 270 850 410,295 ^ csf 2 825 535,480,180 -- -- 300, 270 850 455,400,295 ^ csf 3 825 535,480,180 -- -- 300, 270 850 455,400,295 ^ cly 2 825 535,480,180 -- -- 300, 270 850 455,400,295 ^ cly 3 825 535,480,180 -- -- 300, 270 850 510,460,410,320 ^ cly 4 825 535,480,180 -- -- 300, 270 850 455,400,295 ^ cly 5 825 535,480,180 -- -- 300, 270 850 510,460,410,320 ^ cly 6 825 535,480,180 -- -- 300, 270 850 510,460,410,320 ^ ben 1 825 470,430,240 390,360 -- 150 850 470,390 870, 530, 355, 340 ben 2 825 470,430,240 390,360 -- 150 850 470,390 870, 530, 355, 340 ben 3 825 470,430,240 385 -- 150 850 470,390 870, 530, 355, 340 blk 1 825 470,430,240 390,360 -- 150 850 470,390 870, 530, 355, 340 blk 2 825 470,430,240 385 -- 150 850 470,390 870, 530, 355, 340 cly 7 825 470,430,240 390,360 -- 150 850 470,390 870, 530, 355, 340 hlm 1 825 470,430,240 385 -- 150 850 470,390 870, 530, 355, 340 hlm 2 825 470,430,240 390,360 -- 150 850 470,390 870, 530, 355, 340 hls 1 825 470,430,240 385 -- 150 850 470,390 870, 530, 355, 340 jns 1 825 470,430,240 385 -- 150 850 470,390 870, 530, 355, 340 mlr 1 825 470,430,240 385 -- 150 850 470,390 870, 530, 355, 340 mlr 2 825 470,430,240 390,360 -- 150 850 470,390 870, 530, 355, 340 pch 1 825 470,430,240 385 -- 150 850 470,390 870, 530, 355, 340 pch 2 825 470,430,240 385 -- 150 850 470,390 870, 530, 355, 340

pch 3 825 470,430,240 390,360 -- 150 850 470,390 870, 530, 355, 340 35

Table 3 (cont.):

ISOLATEa LR12R LR12R ATA-1 AME-1 AtITS1 ITS1 ITS1/ ITS1/ O-1 O-1 ATA-2 AME-2 AmITS1 ITS4 ITS4 ITS4 (bp)b Alu I (bp)d (bp)e ITS2 (bp)g Hha I Mbo II (bp)c (bp)f (bp)h (bp)i stc 1 825 470,430,240 390,360 -- 150 850 470,390 870, 530, 355, 340 swl 1 825 470,430,240 390,360 -- 150 850 470,390 870, 530, 355, 340 tms 1 825 470,430,240 385 -- 150 850 470,390 870, 530, 355, 340 tms 2 825 470,430,240 390,360 -- 150 850 470,390 870, 530, 355, 340 tms 3 825 470,430,240 390,360 -- 150 850 470,390 870, 530, 355, 340 ttn 1 825 470,430,240 390,360 -- 150 850 470,390 870, 530, 355, 340 usd 1 825 470,430,240 390,360 -- 150 850 470,390 870, 530, 355, 340 usd 2 825 470,430,240 385 -- 150 850 470,390 870, 530, 355, 340 ctn 1 825 470,430,325,240 390,360 -- 150 850 470,390 870, 530, 355, 340 dky 1 825 470,430,325,240 390,360 -- 150 850 470,390 870, 530, 355, 340 gmg 1 825 470,430,325,240 390,360 -- 150 850 470,390 870, 530, 355, 340 gmg 2 825 470,430,325,240 390,360 -- 150 850 470,390 870, 530, 355, 340 ync 1 825 470,430,325,240 390,360 -- 150 850 520, 215 870, 530, 340 ync 2 825 470,430,325,240 390,360 -- 150 850 520, 215 870, 530, 340 ync 3 825 470,430,325,240 385 -- 150 850 520, 215 870, 530, 340 a Table 1 describes the isolates b Amplicon size following PCR amplification with primer set LR12R/O-1. c Amplicon size following PCR amplification with primer set LR12R/O-1 and restriction with Alu I. d Amplicon size following PCR amplification with primer set ATA-1/ATA-2. e Amplicon size following PCR amplification with primer set AME-1/AME-2. f Amplicon size following PCR amplification with primer set AtITS1/AmITS1/ITS2. g Amplicon size following PCR amplification with primer set ITS1/ITS4. h Amplicon size following PCR amplification with primer set ITS1/ITS4 and restriction with Hha I. i Amplicon size following PCR amplification with primer set ITS1/ITS4 and restriction with Mbo II. -- No amplicon produced. ^ Pattern insufficient to classify isolate. 36

Figure 3: Analysis of the IGS and ITS of known A. tabescens isolate TA1 and known A. mellea isolate ST5. A. TA1 shows a band of 385 bp and ST5 shows no amplification with primer set Ata-1/Ata-2. Neither isolate has a product amplified with Ame-1/Ame-2. With the ITS-1/ITS-4 combination, both isolates demonstrated a band of 850 bp. Primer set AtITS-1/AmITS-1/ITS-2, amplification of ST5 produced a band of 270 bp while TA1 amplified a band of 150 bp. B. After amplification with primer set LR12R/O-1 and restriction with Alu I, ST5 revealed bands of 535, 480, and 180 bp and TA1 showed bands of 430 and 240 bp. C. Amplification with primer set ITS-1/ITS-4 of TA1, followed by restriction with Mbo I produced bands of 870, 530, 355, and 340 bp while Hha I produced bands of 470 and 390 bp.

Figure 4: IGS region of Armillaria isolates amplified with primers LR12R/O-1 and restricted with Alu I. A. PCR amplification produced an amplicon of approximately 825 bp for all isolates. B. Upon digestion with Alu I, three banding patterns emerged. Putative A. mellea isolates exhibited bands approximately 535, 480, and 180 bp. Putative A. tabescens isolates had banding patterns of either 470, 430, and 240 bp (usd1 and pst3) or 470, 430, 325, and 240 bp (gmg2 and ync1).

Figure 5: IGS region of Armillaria isolates amplified with primers based on the sequences of European isolates (Sicoli et al., 2003) to identify A. mellea and A. tabescens. A. Primer set Ata- 1/Ata-2 did not amplify any products from putative A. mellea isolates. Ata-1 and Ata-2 amplified a band of approximately 385 bp in the putative A. tabescens isolates. B. Primer set Ame-1/Ame-2 failed to produce amplicons in any of the isolates.

Figure 6: ITS region of Armillaria isolates amplified with primer set ITS-1/ITS-4 and the primer set developed by Bryson et al. (2003) to discriminate among southeastern Armillaria spp. A. With universal primers ITS-1 and ITS-4 an amplicon of approximately 850 bp was amplified in all of the isolates. B. With primer set AtITS-1, AmITS-1, and ITS-2 (Bryson et al., 2003), most putative A. mellea isolates exhibited a doublet pattern of 300 and 270 bp, while all putative A. tabescens isolates showed a single band of 150 bp.

Figure 7: ITS regions of putative A. tabescens isolates amplified with primer set ITS-1/ITS-4 and restricted with Hha I and Mbo II to visualize polymorphisms among isolates. A. Hha I restriction revealed two groups among the isolates. One group composed of the ync isolates had bands of 520 and 215 bp while all other isolates had bands of 470 and 390 bp. B. Mbo II restriction revealed the same two groups. The ync isolates had bands of 870, 530, and 340 bp while all other isolates had bands of 870, 530, 355, and 340 bp.

Figure 8: ITS regions of putative A. mellea isolates amplified with primer set ITS-1/ITS-4 and restricted with Hha I. Four banding patterns were shown. Isolate csf 1 had a pattern of 500, 470, 400, and 295 bp. Isolates csf 2, csf 3, cly 2, and cly 4 all had patterns of 455, 400, and 295 bp. Isolates cly 3, cly 5, and cly 6 each exhibited bands of 510, 460, 410, and 320 bp. Isolate cly 9 had bands of 410 and 295 bp.

CHAPTER 4

DISCUSSION

With the appearance that Armillaria root rot is becoming an increasing threat to tree health in the southeastern peach industry, the urgency to understand and control the disease becomes increasingly important. Currently, resistant rootstock materials offer the most promise for future control. A greater comprehension of the variability within and among species relating to both pathogenicity and virulence will provide focus to breeding efforts. In order to perform pathogenicity and virulence studies, accurate species and perhaps even below-species classification is necessary.

This work was undertaken to gain a better understanding of the species of Armillaria that infect southeastern peach orchards. Originally, the goals were to differentiate among isolates at a below-specifies level and to establish a better understanding of isolate pathogenicity based on these differences. However, while using the accepted methods of amplification and restriction analysis to classify isolates, it was realized that this group would be more complicated than previously thought. In addition, there were difficulties and discrepancies with conventional mating studies, as well as with DNA analysis.

DNA extraction. Initially, attempts were made to directly amplify DNA from small quantities of Armillaria mycelia (Harrington and Wingfield, 1995). The use of this method was desirable since it did not require DNA extraction prior to amplification. It became apparent that this method was not going to be as convenient as previously described. Direct amplification resulted in DNA of poor quality that gave inconsistent and poor PCR amplifications. In the 20% of reactions that did amplify, the PCR product appeared to be either of low quality or the reaction

44 had an inadequate amount of material. Although direct amplification seemed quicker and easier, in the long run it would require multiple attempts for each isolate in order to achieve successful reactions, leaving doubt as to the validity of the results because of poor PCR products.

Therefore, other DNA extraction techniques were explored.

After the low success rate of direct amplification, other techniques to extract fungal DNA were attempted (Lee and Taylor, 1990). The Lee and Taylor method produced a higher yield of amplifiable DNA most of the time. However, poor PCR amplification occurred with the DNA extracted by this method. Large amounts of apparent polyphenolic compounds were observed in the extractant. Thus, it seemed likely that the polyphenolics produced by the fungi were reducing the quality of products of the PCR amplification. Modification of the procedure would be necessary to ‘protect’ the DNA. After exploring DNA extraction techniques of other high phenol-producing tissues, such as cotton, PVP was incorporated into the lysis buffer to remove polyphenolics (John, 1992; Pich and Schubert, 1993). Because PVP removed the phenolics from the DNA there was less interference during PCR.

The challenge was compounded by the appearance of a mite infestation in the incubator.

Therefore, isolates were grown in MYE liquid culture for DNA extraction. The continuation in liquid culture allowed rescue of the cultures preventing mite contamination from destroying the collection. It was ideal to harvest the tissue before the exudates blackened the liquid media.

Apparently the older tissues were saturated with phenolics making it more difficult to obtain pure and intact DNA.

Because small quantities of material (300 mg) were needed to begin DNA extraction, younger, actively growing mycelia on the periphery of MYEA cultures was utilized. DNA extraction from plated cultures required less time and produced intact, pure DNA at quantities

45 comparable to those reported by John (1992) and Pich and Schubert (1993). The tissue was grown in a shorter amount of time (several weeks for agar culture vs. several months for liquid culture) and the only preparation necessary was plate inoculation.

Molecular techniques: whole-genome, IGS, and ITS analysis. Initially, the goal was to develop primers which identified isolates at a species or below-species level by widespread screening with RAPD primers. RAPD primer screening was reported to discriminate among genera, species, and even races (Crowhurst et al., 1991; Shi et al., 1995; Tommerup et al., 1995;

Voight et al., 1998). Because there were so many bands amplified with no obvious repeatable patterns, other approaches were necessary to first characterize the isolates. Apparent morphological definitions of species were inadequate, requiring other forms of isolate identification before any final classification could occur. It was determined that this method of developing species-specific primer sets would be more useful after isolates were identified by more conventional means. Furthermore, the RAPD method may prove more useful with amplified rDNA from ITS or IGS regions.

Harrington and Wingfield (1995) first amplified the IGS region with primers LR12R and

O-1 prior to restriction with various enzymes. They discovered that Alu I was the most useful enzyme to differentiate among Armillaria species, including A. tabescens and A. mellea. After restricting the amplicons obtained from unknown isolates, the patterns were indeed similar enough to those of Harrington and Wingfield (1995) to draw some inferences. The main discrepancy lies in the presence of a ‘doublet’ banding pattern seen in most of the higher molecular weight bands. There are several possible explanations for the presence of this extra band. First, this may be a result of the diploid nature of the fungus. DNA extracted from cultures started from single-spore isolates would be haploid and therefore exhibit only a single

46 set of alleles for this region. Second, the decision to report particular bands is up to the discretion of the investigator. One might choose to report either highly contrasted bands or every single band visualized on the gel regardless of intensity or size. There is evidence of faint banding for some A. tabescens and A. mellea isolates in the Harrington and Wingfield (1995) results, which could be interpreted as doublets, similar to those identified in the results reported here.

Harrington and Wingfield (1995) referred to a group of isolates which produced 430 and

240 bp bands as ‘A. tabescens A’. Similarly, this study revealed that 23 of the southeastern isolates presented bands at 470, 430, and 240 bp. The 470/430 doublet in the southeastern set was consistent with the 430 bp band identified in the ‘A. tabescens A’ by Harrington and

Wingfield (1995). Furthermore, TA1, known to be A. tabescens (a Florida isolate), gave single bands at 430 and 240 bp. TA1 was labeled as a haploid tester strain (Guido Schnabel, personal communication), lending support to the suggestion that doublets are a result of a diploid genome.

Seven of the A. tabescens isolates were similar possessing the same pattern as group I with the addition of an intermediate band at 325 bp. The Harrington and Wingfield (1995) study reports

‘A. tabescens B’ as having a band at 320 bp as well as 240 and 100 bp. While not identical, there was some similarity with our results. In addition, their figure of the ‘A. tabescens B’ Alu I restriction contains a faint band at approximately 420 bp when analyzed with Gel-Pro software.

Both geographic regions (Piedmont and Ridge) as well as all three states (Georgia, South

Carolina, and Alabama) were represented by each sub-grouping of A. tabescens isolates.

A third group was also determined by the Alu I digestion of the IGS region. This group contained ST5 as well as nine isolates collected from the cly (Piedmont) and csf (Ridge) sites in

South Carolina. This group, similar to Harrington and Wingfield’s ‘A. mellea A’ (490 and 180

47 bp), exhibited bands of approximately 535, 480, and 180 bp. Gel-Pro analysis of the Harrington and Wingfield (1995) results indicates a faint band at 530 as well as bands at 490 and 180 bp, similar to our Alu I restriction of A. mellea isolates.

Based upon Alu I data, 30 of the isolates were classified as A. tabescens (Table 4). This set of species can be divided into two groups based on polymorphisms within the IGS region.

‘Group I’ was previously described by Harrington and Wingfield (1995) as ‘A. tabescens A’ and

‘Group II’ has some similarities to their ‘A. tabescens B’ (Table 4). The A. mellea isolates consisted of only one group to which ‘Group I’ was assigned (Table 5). This group is thought to be the same as Harrington and Wingfield’s ‘A. mellea A’. However, because disparities still existed between these Alu I restriction patterns and those reported by Harrington and Wingfield

(1995), it was necessary to explore additional methods of isolate characterization.

Since comparable Alu I restriction patterns were found among Armillaria spp. from North

America and Europe (Harrington and Wingfield, 1995), it was conceivable that primers based on the IGS region of European isolates might be useful in determining the species of these southeastern isolates. Sicoli et al. (2003) designed primer sets to test for A. tabescens (Ata-1 and Ata-2) and A. mellea (Ame-1 and Ame-2).

Upon amplification with the primer set Ata-1 and Ata-2, 30 of the A. tabescens isolates, as well as TA1, produced either a single or double band. Since these primers were developed from the IGS region of A. tabescens and should amplify only this species, this data is taken as further evidence of the identification of the A. tabescens isolates (Table 4). However, this band size was viewed as different from the 330 bp band reported by Sicoli et al. (2003), suggesting that more heterogeneity existed between the European and North American isolates than initially thought. Since this primer set was designed to identify A. tabescens isolates, it was not expected

48 to amplify any of the A. mellea species, nor did it (Table 5). Conversely, the Ame-1 and Ame-2 primers, which should have detected A. mellea failed to produce amplicons in any of the southeastern isolates, including ST5 and TA1. While the Ata-1 and Ata-2 primer set supported the conclusion that the putative A. tabescens isolates were in fact A. tabescens, the Ame-1 and

Ame-2 set did not support a similar conclusion for the A. mellea isolates. Therefore, further investigation was required.

The reportedly less heterogeneous ITS region offered further opportunity to characterize the southeastern isolates in this study. Using primers AtITS-1, AmITS-1 developed by Bryson et al. (2003) and the universal primer ITS-2, the isolates were further classified. Again, the A. tabescens isolates behaved as expected, producing a band of approximately 150 bp. While the A. mellea isolates exhibited a band of approximately 300 bp, in seven out of the nine isolates this band appeared as a doublet, similar to the Alu I results in this study. This doublet was also attributed to multiple alleles.

Restriction with Mbo II and Hha I illustrated that the ITS regions were potentially more hetergeneous than the IGS region when RFLPs were being considered. Armillaria tabescens was divided into two groups when restricted with Hha I and Mbo II. In each case, ‘Group B’ consisted of isolates from the ync location in South Carolina. All other A. tabescens from

Georgia, South Carolina, and Alabama were placed in ‘Group A’ by this method (Table 4).

The group of nine A. mellea isolates was found to be more diverse. Four distinct polymorphic patterns were shown when these isolates were restricted with Hha I (Table 5).

‘Group A’ and ‘Group C’ were represented at the csf site, while ‘Group B’, ‘Group C’, and

‘Group D’ were represented at the cly site. The known A. mellea, ST5, did not reveal any pattern

49 with Hha I, nor did any putative A. mellea isolate among the unknowns assessed in this study with Mbo II.

The Mbo II and Hha I restriction patterns were different from those predicted based on the European ITS sequences obtained from the NCBI database. The only similarities seen between the predicted and actual results are that two cleavage sites were made when A. tabescens was restricted with Hha I. However, the band sizes were inconsistent with predicted sizes.

Thus, European and North American isolates may be quite different.

The diversity within such a limited number of A. mellea isolates is interesting. Not only is there a wide range of heterogeneity within the species, there is also a substantial diversity within a given location. Based on this restriction analysis, three genotypes were found at the cly location and two were found at the csf location in South Carolina. This raises the questions as to the extent of hydridization and recombination that may be occurring among these A. mellea isolates to cause varying polymorphisms within the highly conserved ITS region.

Conclusion

This research was undertaken to gain a better understanding of the Armillaria spp. that affect southeastern peach orchards. This knowledge is important in that it will not only lead to an increased understanding of the variability of these isolates, but also lend focus to resistance breeding programs. Ultimately, it will be helpful to assess the diversity of pathogenicity and virulence of this group of fungi as control methods are being researched.

Although attempts at morphological techniques and molecular analysis using RAPD primers proved unfruitful, they did strengthen the understanding that this group possessed a great degree of diversity. It was surmised that development of unique primers using a RAPD approach

50 would hold more merit once the species of the unknown isolates were determined using established techniques. The IGS and ITS regions of the rDNA were then explored as a means of identification. Through an array of primers and protocols cited in previous work (Bryson et al.,

2003; Harrington and Wingfield, 1995; Sicoli et al., 2003), the unknown isolates of this southeastern collection were characterized. The Bryson et al. (2003) method was preferred above the other techniques for determining Armillaria spp. This method required only PCR amplification of the ITS-1 region with no subsequent restrictions. However, if below-species classification was sought, one of the IGS or ITS amplification and restriction analyses was required.

Many times there were slight discrepancies between the results of this study and previously reported data. However, there are several possible explanations for these differences.

By using amplification and restriction of the IGS and ITS regions, the group of 39 southeastern isolates has been divided into 30 isolates of A. tabescens and nine of A. mellea. Although the isolate collection was not obtained using a random sampling approach, this research supports the idea that A. tabescens is the primary species involved with decline in southeastern peach orchards. However, A. mellea was also found in two South Carolina orchards. This research therefore suggests that A. mellea is not as widespread in southeastern peach orchards as A. tabescens.

Further classification was also possible by restricting the IGS region with Alu I. Based on the RFLP analysis, two groups of A. tabescens and one group of A. mellea were identified.

Conversely, when the ITS region was restricted with Mbo II and Hha I, it suggested that there were two groups present within A. tabescens and four within A. mellea. It was surprising that four polymorphisms existed within the nine A. mellea isolates, especially since the ITS region is

51 considered to be more conserved than the IGS region. This finding suggests that there is high degree of heterogeneity within this region of the A. mellea genome.

When ITS and IGS are considered together to characterize this group of isolates, three major ‘groups’ have emerged within A. tabescens (Table 4). Group 1 consists of isolates which belong to IGS ‘group I’ and ITS ‘group A’. Group 2 is composed of isolates belonging to IGS

‘group II’ and ITS ‘group A’. Group 3 contains isolates from IGS ‘group II’ and ITS ‘group B’.

When a similar analysis was done with the A. mellea isolates, four major ‘groups’ emerge;

‘group 4’, ‘group 5’, ‘group 6’, and ‘group 7’ (Table 5). When analyzing the geographical distribution of the seven ‘groups’ established in this study, the most noticeable trend was that the

A. mellea isolates were found only in the northernmost counties of South Carolina (Figure 9). In addition, ‘Group 1’ of the A. tabescens isolates appeared to be the most widely distributed and

‘group 3’ was found in only one county (Figure 9).

While this research extended molecular characterizations of the southeastern isolates, much work remains. A more comprehensive screening should be conducted to reveal any other potential patterns that exist in the current collection as well as additional areas throughout the southeastern peach region. Potential research will require DNA sequences of the ITS and IGS regions of the representative members of each group to identify genomic differences among the isolates. To aid in directing the focus of resistance breeding, pathogenicity analyses may be implemented with representative isolates in each group.

Table 4: Identification and classification of southeastern Armillaria tabescens isolates.

IGS ITS Alu I Alu I ATA-1 AME-1 ITS1 Hha I Mbo II IGS ATA-2 AME-2 AmITS1 + ITS2 ITS Isolatea Species Groupb Species Speciesc Speciesd Group Group Group TA1 A. tab. I A. tab. -- A. tab. A A 1 ben 1 A. tab I A. tab. -- A. tab. A A 1 ben 2 A. tab I A. tab. -- A. tab. A A 1 ben 3 A. tab I A. tab. -- A. tab. A A 1 blk 1 A. tab I A. tab. -- A. tab. A A 1 blk 2 A. tab I A. tab. -- A. tab. A A 1 cly 7 A. tab I A. tab. -- A. tab. A A 1 hlm 1 A. tab I A. tab. -- A. tab. A A 1 hlm 2 A. tab I A. tab. -- A. tab. A A 1 hls 1 A. tab I A. tab. -- A. tab. A A 1 jns 1 A. tab I A. tab. -- A. tab. A A 1 mlr 1 A. tab I A. tab. -- A. tab. A A 1 mlr 2 A. tab I A. tab. -- A. tab. A A 1 pch 1 A. tab I A. tab. -- A. tab. A A 1 pch 2 A. tab I A. tab. -- A. tab. A A 1 pch 3 A. tab I A. tab. -- A. tab. A A 1 stc 1 A. tab I A. tab. -- A. tab. A A 1 swl 1 A. tab I A. tab. -- A. tab. A A 1 tms 1 A. tab I A. tab. -- A. tab. A A 1 tms 2 A. tab I A. tab. -- A. tab. A A 1 tms 3 A. tab I A. tab. -- A. tab. A A 1 ttn 1 A. tab I A. tab. -- A. tab. A A 1 usd 1 A. tab I A. tab. -- A. tab. A A 1 usd 2 A. tab I A. tab. -- A. tab. A A 1 ctn 1 A. tab II A. tab. -- A. tab. A A 2 dky 1 A. tab II A. tab. -- A. tab. A A 2 gmg 1 A. tab II A. tab. -- A. tab. A A 2 gmg 2 A. tab II A. tab. -- A. tab. A A 2 ync 1 A. tab II A. tab. -- A. tab. B B 3 ync 2 A. tab II A. tab. -- A. tab. B B 3 ync 3 A. tab II A. tab. -- A. tab. B B 3 a Table 1 describes isolates b Similar to patterns reported by Harrington and Wingfield (1995) c No amplicon produced d Similar to band size reported by Bryson et al. (2003)

Table 5: Identification and classification of southeastern Armillaria mellea isolates.

IGS ITS Alu I Alu I ATA-1 AME-1 ITS1 Hha I Mbo II IGS ATA-2 AME-2 AmITS1 + ITS2 ITS Isolatea Species Group b Speciesc Speciesc Speciesd Group Groupe Group ST5 A. mel. I -- -- A. mel. *e * csf 1 A. mel. I -- -- A. mel. A * 4 cly 1 A. mel. I -- -- A. mel. B * 5 csf 2 A. mel. I -- -- A. mel. C * 6 csf 3 A. mel. I -- -- A. mel. C * 6 cly 2 A. mel. I -- -- A. mel. C * 6 cly 3 A. mel. I -- -- A. mel. D * 7 cly 4 A. mel. I -- -- A. mel. C * 6 cly 5 A. mel. I -- -- A. mel. D * 7 cly 6 A. mel. I -- -- A. mel. D * 7 a Table 1 describes the isolates b Similar to pattern reported by Harrington and Wingfield (1995) c No amplicon produced d Similar to band size reported by Bryson et al. (2003) e Inconclusive results

Code County Isolates IGS/ITS Groupa AL-1 Chilton ctn 1 2 swl 1 1 GA-1 Oconee tms1, tms 2, tms 3 1, 1, 1 GA-2 Crawford dky 1 2 GA-3 Peach usd 1, usd 2 1, 1 SC-1 Spartanburg cly 1, cly 2, cly 3, 5, 6, 7 cly 4, cly 5, cly 6, cly 7 6, 7, 7, 1 gmg 1, gmg 2 2, 2 SC-2 York ben 1, ben 2, ben 3 1, 1, 1 blk 1, blk 2 1, 1 mlr, 1, mlr 2 1, 1 pch 1, pch 2, pch 3 1, 1, 1 SC-3 Chesterfield csf 1, csf 2, csf 3 4, 6, 6 SC-4 Saluda Hls 1 1 Ttn 1 1 SC-5 Edgefield hlm1, hlm 2 1, 1 jns 1 1 stc 1 1 ync 1, ync 2, ync 3 3, 3, 3

a Groups established by IGS and ITS molecular analysis. Summary of groups in Table 4 and Table 5.

Figure 9: Geographical distribution of Armillaria “groups” as established by molecular characterization of IGS and ITS regions.

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