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2007 Epizootiology and Phylogenetics of Entomopathogenic Fungi Associated with Fiorinia externa ferris(Hemiptera: Diaspididae) in the Northeastern USA Jose A. P. Marcelino University of Vermont

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Recommended Citation Marcelino, Jose A. P., "Epizootiology and Phylogenetics of Entomopathogenic Fungi Associated with Fiorinia externa ferris(Hemiptera: Diaspididae) in the Northeastern USA" (2007). Graduate College Dissertations and Theses. 148. https://scholarworks.uvm.edu/graddis/148

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EPIZOOTIOLOGY AND PHYLOGENETICS OF ENTOMOPATHOGENIC

FUNGI ASSOCIATED WITH FIORINIA EXTERNA FERRIS (HEMIPTERA:

DIASPIDIDAE) IN THE NORTHEASTERN USA

A Dissertation Presented

José A. P. Marcelino

The Faculty of the Graduate College

The University of Vermont

In Partial fulfillment of the Requirements for the Degree of Doctor of Philosophy Specializing in Insect Pathology

October, 2007 Accepted by the Faculty of the Graduate College, The University of Vermont, in partial fulfillment of the requirements for the degree of Doctor of Philosophy, specializing in Plant and Soil Sciences.

Date: August 24th, 2007

1 Abstract

The eastern hemlock [Tsuga canadensis (L.) Carrière] is one of the native dominant forest components of northeastern US. At present, these valuable stands face an alarming decline, in part due to the Fiorinia externa, elongate hemlock scale (EHS), (Hemiptera: Coccoidea: Diaspididae). The armored shield of F. externa provides an excellent defense against insecticides, natural enemies and adverse conditions. Chemical and classical biocontrol methods have been unable to stop the spread of this pest. Recently, the occurrence of an epizootic within the F. externa population in the Mianus River Gorge Preserve in Bedford, NY revealed a promising opportunity for control of this scale. Entomopathogenic fungi represent a valuable, although under-utilized, group of organisms with unique capabilities for self-sustaining pest management. Given the significant impact of this epizootic on F. externa, we have conducted extensive research on the biology, genetics and biological control potential of this epizootic.

We molecularly identified a complex of entomopathogenic, phytopathogenic, and endophytic fungi associated with the epizootic in 36 localities within the states of New York, Pennsylvania, Connecticut and New Jersey. One fungus, Colletotrichum sp., was the most commonly isolated organism in populations of F. externa within areas of the epizootic. The host range of this Colletotrichum species comprised both insects and plants, although diverse life cycles occured in the different hosts. Endophytic growth was observed in 28 species of plants comprising 18 families (52% of the sampling), whereas in F. externa biotrophic and necotrophic growth was detected. Colletotrichum is a widely known phytopathogenic genus and reports of entomopathogenic activity are extremely rare. In order to understand the biological processes involved in the host-pathogen interactions we quantified the pathogenicity and virulence of this Colletotrichum sp. to four insect families and six plants families as well as the occurrence of sexual recombination in this Colletotrichum sp., both in vitro and in planta.

We observed that this Colletotrichum sp. displays a propensy to induce rapid disease and mortality in F. externa hosts. Phylogenetic analysis comprising six of the most commonly studied nuclear genes in molecular phylogenetics (D1/D2 domain of the 28 rDNA gene, ITS region, β-Tubulin 2, GPDH gene, GS gene and HMG box at the MAT1-2 mating-type gene) and RAPDs showed this fungus is closely related to phytopathogenic strains of Colletotrichum acutatum and that it may represent a single population lineage of this species (i.e., Colletotrichum acutatum forma specialis fiorinia). Though a large body of information exists regarding the phytopathogenic genus Colletotrichum, ours is only the second reported entomopathogenic strain. It is not clear whether the colonization of an insect by this fungus is truly rare or a common but undetected event. Sexual recombination, observed in planta and in vitro, could be the means by which new genetic variants are generated leading to new biotypes with a selective advantage to colonize new hosts, which in this case is a novel host in a different kingdom.

2 Citations

Material from this dissertation has been submitted for publication to Forest Science, Mycologia and Journal of Insect Science on September 15th 2007 in the following form:

José A. P. Marcelino, Svetlana Gouli, Rosanna Giordano, Vladimir V. Gouli, Bruce L. Parker, Margaret Skinner. Fungi associated with a natural epizootic in Fiorinia externa Ferris (Hemiptera: Diaspididae) populations. Forest Science.

Marcelino J.A.P., Giordano R., Gouli S., Gouli, V. V., Parker B. L., Skinner, M., Cesnik R., TeBeest D. Colletotrichum acutatum f. sp. fiorinia infection of a scale insect, Fiorinia externa Ferris (Hemiptera: Diaspididae). Mycologia.

José A. P. Marcelino, Svetlana Gouli, Bruce L. Parker, Margaret Skinner, Rosanna Giordano, Lora Schwarzberg. Host Plant Associations with an Entomopathogenic Strain of Colletotrichum acutatum from Elongate Hemlock Scale. J. Insect Science.

José A. P. Marcelino, Svetlana Gouli, Vladimir V. Gouli, Bruce L. Parker, Margaret Skinner. Entomopathogenic Activity of a Colletotrichum acutatum Strain Recovered from Elongate Hemlock Scale. J. Insect Science.

Acknowledgments

I am very grateful to a number of people for their assistance and encouragement during

my studies at the University of Vermont, to whom I feel indebt.

I would like to thank my supervisor, Dr. Bruce L. Parker, for his guidance in this project

and to Drs. Svetlana Gouli and Rosanna Giordano for sharing their vast knowledge in fungi and genetics with me, as well as, for their kind friendship and perseverance (and

food!). I would also like to thank the valuable ideas and suggestions given by Drs.

Margaret Skinner, Vladimir Gouli, David TeBeest, Felipe Neri Soto-Adames, Brent

Teillon and Dale Bergdahl during the course of this project. I must thank as well all the

staff of the Entomology Research Laboratory and the Plant and Soil Science Department,

for their help and valuable technical assistance and Mr. Jon Turmel, for being an example

of what it means to be an excelent teacher. Also, I have to thank Lora Schwarzberg,

which was instrumental in achieving some of the goals in this work.

My task would have not been possible without their help and professional expertise.

I would also like to thank the encouragement and support of my friends. Luisa, Dina,

Duarte, Felipe and Carla who, from a distance, lived through much of my experiences

during this journey, and to my roommate Trevor which patiently listen to me every day.

Also to Ana Melo, which watched me from the distance and then from the stars.

Finally I would like to thank the unique opportunity that the University of Vermont has

given me to grow as a professional and as a person in the US.

iii

Table of Contents

Citations ...... ii Acknowledgments...... iii List of Tables ...... vi List of Figures...... vii

CHAPTER 1. LITERATURE REVIEW ...... 2

1. Introduction...... 2 1.1. Importance and decline of eastern hemlock...... 4 1.2. The Fiorinia externa (Hemiptera: Coccoidea: Diaspididae) armored scale...... 9 1.2.1. Life-cycle ...... 12 1.2.2. Damage ...... 14 1.2.3. Management...... 15 1.2.3.1. Chemical control...... 16 1.2.3.2. Natural enemies: parasites and predators...... 17 1.2.3.3. Entomopathogenic fungi...... 18 1.3. Entomopathogenic fungi as biological control agents...... 21 1.4. The genus Colletotrichum...... 26 1.4.1. Taxonomic considerations ...... 27 1.4.2. rDNA analyses to discern Colletotrichum spp...... 30 1.4.3. Hosts ...... 32 1.4.3.1. Plants...... 32 1.4.3.2. Mammals and amphibians ...... 36 1.4.3.3. Insects ...... 37 1.4.4. Biocontrol of weeds with Colletotrichum spp...... 38 1.4.5. Mating systems in Colletotrichum spp...... 40 1.4.6. Recombinant Colletotrichum spp...... 42 1.4.7. Infection process ...... 44 1.4.8. Quiescent infection by Colletotrichum spp...... 46 1.4.9. Survival and dispersal under different abiotic conditions...... 48 1.5. Research objectives...... 50

CHAPTER 2. FUNGI ASSOCIATED WITH A NATURAL EPIZOOTIC IN FIORINIA EXTERNA

FERRIS (HEMIPTERA: DIASPIDIDAE) POPULATIONS...... 58

CHAPTER 3. COLLETOTRICHUM ACUTATUM F. SP. FIORINIA INFECTION OF A SCALE INSECT,

FIORINIA EXTERNA FERRIS (HEMIPTERA: COCCOIDEA) ...... 76

CHAPTER 4. HOST PLANT ASSOCIATIONS WITH AN ENTOMOPATHOGENIC STRAIN OF

COLLETOTRICHUM ACUTATUM FROM ELONGATE HEMLOCK SCALE ...... 120

CHAPTER 5. ENTOMOPATHOGENIC ACTIVITY OF A COLLETOTRICHUM ACUTATUM STRAIN

RECOVERED FROM ELONGATE HEMLOCK SCALE ...... 144

CHAPTER 6. COMPREHENSIVE BIBLIOGRAPHY ...... 162

APPENDICES...... 195

Appendix 1. The genus Aschersonia ...... 195

List of Tables

Table 1.1. Reports of fungal species recovered from populations of scale insects...... 51 Table 1.2. Reports of fungi recovered from Fiorinia species ...... 56 Table 1.3. Sequence of morphological identifications of the most commonly isolated fungus from diseased F. externa in the states of Connecticut, New York, New Jersey and Pennsylvania ...... 57 Table 2.1. Pure fungal isolates recovered from Fiorinia externa ...... 63 Table 3.1. Characterization of different Colletotrichum spp...... 111 Table 3.2. Morphological plasticity in Colletotrichum sp. isolated in epizootic areas.. 112 Table 3.3. Cross-fertile bioassay in toothpick substratum ...... 113 Table 3.4. Conidia morphology of entomopathogenic Colletotrichum spp. at 22 C in PDA medium examined in water and stained with cotton blue...... 114 Table 3.5. Appressoria morphology of entomopathogenic Colletotrichum sp. strains growing in strawberry leaves at 22 C examined in water and stained with cotton blue. 116 Table 3.6. Perithecia of Glomerella sp. on snap bean stems (2 cm) at 22 C from cross- fertile ERL1385 x EHS58 ...... 118 Table 3.7. Ascospores of Glomerella sp. on snap bean stems (2 cm) at 22 C and under natural light ...... 119 Table 4.1. Fungal isolates used for the horticultural plant and hemlock phenological bioassays ...... 123 Table 4.2. Percentage of fungal re-isolations obtained from the test plants (N = 8 plants /isolate and plant species...... 130 Table 4.3. Percentage of eastern hemlock seedlings from which Colletotrichum sp. was re-isolated after treatment (N = 4 seedlings/age and month)...... 133 Table 4.4. Plant material sampled in epizootic areas and screened for C. acutatum (a sample of live leaves from one plant per species was taken unless otherwise indicated) ...... 135 Table 5.1. Fungal isolates tested in the insect bioassays...... 147 Table 5.2. Results of statistical analysis for bioassays with Fiorinia externa immatures ...... 154 Table 5.3. Statistical analysis for bioassays with Spodoptera exigua, Bemisia argentifolii and Frankliniella occidentalis ...... 155

List of Figures

Figure 1.1. Distribution of hemlock woolly adelgid, Adelges tsugae, in the eastern US .. 7 Figure 1.2. Distribution of elongate hemlock scale, Fiorinia externa, in the eastern US . 8 Figure 1.3. F. externa adult female carrying eggs under translucent exuvia (A) and F. externa adult male with profuse filamentous waxy strands (B) ...... 10 Figure 1.4. F. externa crawler...... 11 Figure 1.5. Settled F. externa nymph secreting glutinose substance from Malphigian tubules and you adult female with full developed fresh scale cover ...... 12 Figure 1.6. Parasitized scale with Encarsia sp. (A) and single Encarsia sp. parasitoid (B) ...... 18 Figure 1.7. F. externa imagoe presenting sclerotia masses ...... 19 Figure 1.8. Taxonomic classification of entomopathogenic fungi...... 24 Figure 2.1. Geographical distribution of sites with diseased Fiorinia externa populations (2002-2006)...... 61 Figure 2.2. Fiorinia externa showing typical symptoms of fungal infection, and representative propagules from several entomopathogenic fungi found associated with F. externa in epizootic areas...... 67 Figure 3.1. Perithecia in planta from cross fertile strains of Colletotrichum sp., ERL1385xEHS58 (A); intact ascus (B); free ascospores (C); Perithecia discharging ascospores (D); appressoria of Colletotrichum sp., strain EHS47, from epizootic site (E); appressorium of C. gloeosporioides, strain EMA26, from Brazilian epizootic (F); Conidia from C. gloeosporioides, EMA26 (G) and ARSEF4630 (H); Colletotrichum sp., strain EHS48, from epizootic area (I); morphological plasticity and overlapping phenotypes of Colletotrichum sp. strains from epizootic localities (J-M); Top (N) and reverse (O) of

Colletotrichum sp. strain EHS58 in PDA at 25 C; Conidia produced by Colletotrichum sp. from epizootic site (P); Strawberry stem from cross fertile bioassay presenting profuse conidia masses (Q); Birch toothpick from cross fertile bioassay with conidia masses and sterile tri-dimensional structures (R); Adult O. praelonga infected with C. gloeosporioides f. sp. ortheziidae (orange masses), Brazil (S); Adult F. externa with epizootic sclerotia masses (T)...... 105 Figure 3.2. Random Amplified Polymorphic DNA (RAPDs) for Colletotrichum sp. strains from different geographic localities within the epizootic area...... 106 Figure 3.3. Phylogenetic tree (Inferred MP tree) among Colletotrichum spp. and Colletotrichum sp. strains isolated from F. externa, for the ITS region...... 107 Figure 3.4. Phylogenetic tree (Inferred MP tree) among Colletotrichum spp. and Colletotrichum sp. strains isolated from F. externa, for the GS gene...... 108 Figure 3.5. Phylogenetic tree (Inferred MP tree) among Colletotrichum spp. and Colletotrichum sp. strains isolated from F. externa, for the GPDH gene...... 109

vii

Figure 3.6. Phylogenetic tree (Inferred MP tree) among Colletotrichum spp. and Colletotrichum sp. strains isolated from F. externa, for the High Mobility Box at the MAT 1-2 gene...... 110

Figure 4.1. Endophytic growth of Fiorinia strain EHS48 in snap bean stem 24h after inoculation, in vitro, showing germinated germ tubes (a) and appressoria (b); (B) Strawberry with necrotic lesions, 2 mo after spraying with the Fiorinia strain ...... 129 Figure 4.2. Adjusted odds ratios (95% confidence interval) of endophytic re-isolations obtained with each fungal inoculation treatment in the plant bioassays...... 131 Figure 4.3. Adjusted odds ratios (95% confidence interval) of endophytic re-isolation obtained for each plant tested in the plant bioassays ...... 132 Figure 4.4. Adjusted odds ratios (95% confidence interval) of endophytic reisolation obtained with fungal treatments in the phenological bioassays...... 134 Figure 5.1. Fungal structures from bioassays of the Fiorinia strain and symptomatic and non-symptomatic F. externa ...... 153 Figure 5.2. Percent mortality among F. externa settlers and crawlers in fungal bioassays. (± 95% CI of the mean) ...... 156 Figure 5.3. Percent mortality among different test insects in fungal bioassays. (± 95% CI of the mean) ...... 157

viii

To my family with deep gratitude

“In wilderness is the preservation of the world” – Thoreau

Published in House & Garden and reprinted with permission of Martyn Thompson©

Sculpture by Antony Gormley, Sitka spruce forest, Scotland. CHAPTER 1

Literature review

1. Introduction

The health of the eastern hemlock forests [Tsuga canadensis (L.) Carr.] in the northeastern US is in decline. Elongate hemlock scale (EHS), Fiorinia externa Ferris

(Hemiptera: Coccoidea: Diaspididae) and hemlock woolly adelgid (HWA), Adelges tsugae Annand (Hemiptera: Adelgidae), have been identified as primary causes in the decline. Initial observations suggested that as A. tsugae invaded stands previously occupied by F. externa, the latter species could be displaced (McClure, 1997). However, the rapid spread of F. externa within the area of A. tsugae seems to contradict this assumption. Strong correlations have been found between A. tsugae and scale infestation levels (Danoff-Burg and Bird, 2002). F. externa can be more of a problem, even when relegated to a secondary role to A. tsugae since this species possesses a high fertility rate as well as an effortless mobility to new potential infestation spots by wind (transportation of crawlers up to 100 meters) and other mechanical agents. The unique shield-like cover or armor of this Diaspidid provides protection for the eggs and the adult from contact insecticides, natural enemies and adverse conditions. Because of its high reproductive rate, losses are soon replaced even with mortality rates of 95% (Baranyovits, 1953,

Johnson and Lyon, 1988). The dense hemlock canopy makes complete coverage with pesticides difficult to achive. This protection is enhanced because most of the scales occur in the lower part of hemlock needles. Systemic insecticides, such as Imidacropid, have also not been effective in controlling the pest (Hoover, 2003). Introduced predators

and parasitoids have not attained significant control of F. externa due to asynchrony

between the natural enemies and pest. F. externa has become a destructive pest of

hemlocks and as evidence of hemlock decline becomes more pronounced and

widespread, public awareness and concern for the potential impact of this species has

grown:

“While the hemlock woolly adelgid gets more of the publicity, other insects are contributing to the decline of the eastern hemlock. Fiorinia externa has probably caused more decline of hemlocks in terms of quick death than the Adelges tsugae”. (G. Hoover, pers. comm., 2004)

“Advanced stage of hemlock decline, as indicated by needle loss, was significantly correlated with Fiorinia externa abundance but not Adelges tsugae abundance, in eastern hemlock stands in the Black Rock Forest, Orange County, New York”. (Danoff-Burg and

Bird, 2002)

“In recent years the populations of F. externa have increased dramatically and have contributed significantly to hemlock decline throughout much of the A. tsugae infested area. In the last 20 years F. externa has significantly expanded its geographical range”.

(McClure, 2002a)

As hemlock declines in vigor, an open canopy allows an increase in understory light levels. As a result, hardwood species especially black birch (Betula lenta L.)

3 establish, leading to major landscape changes by transforming large areas dominated by conifers to young, rapidly growing hardwoods (Foster and O’Keefe, 2000). Moderate winter temperatures have favored the survival of F. externa and A. tsugae and populations of both species have increased. If dispersal of F. externa continues unimpeded and at the present rate, drastic changes in forest composition, landscape and ecological balance can be foreseen and ultimately the disappearance of the current hemlock stands and its historical, economical, environmental protective and recreational value could be lost.

A disquieting resemblance with chestnut stands (Castanea dentata (Marshal)

Borkh.) in the early 1900s can be observed. The rapid spread of chestnut blight

[Cryphonectria parasitica (Murrill) Barr] dictated the fate of the species and practically eradicated 3.5 billion chestnut trees from eastern North America in the following 40 years

(Roane et al., 1986; Anagnostakis, 1988; Bisiach, 1992).

1.1. Importance and decline of eastern hemlock

The eastern or Canada hemlock [Tsuga canadensis (L.) Carrière] is a native species of the eastern US and adjacent Canada. It is one of the more low light tolerant trees in New England forests (Burns, 1923; Anderson and Gordon, 1994) and one of the most long-lived in the eastern US, living up to 300 years (Godman and Lancaster, 1990;

Foster, 2000). Eastern hemlock has been a dominant component of the New England forests for 8000 yr (Foster and Zebryk, 1993). The Pleistocene glaciations eliminated vegetation in the northeastern US and Canada, reducing hemlock to more moderate climatic conditions along the coast or in the foothills of the Appalachians (Anderson and

Gordon, 1994). Pollen records show dramatic fluctuations of eastern hemlock in

northeastern forests suggesting that this species recovered after each fluctuation, regaining dominance in certain areas (Foster and Zebryk, 1993).

Eastern hemlock plays a key role in maintaining balanced, stable forest ecosystems. Their leaf litter produces a forest floor dominated by ammonium nitrogen, as opposed to the nitrate-based nitrogen of deciduous species, thus influencing long-term site quality and species dynamics (Anderson and Gordon, 1994; Mladenoff, 1995). They play a particular important ecological role along streams, where their shade provides shelter, sustains aquatic ecosystems and creates unique microclimate for forest wildlife

(Howe and Mossman, 1995; Wydeven and Hay, 1995; Crow, 1995; Howard et al., 2000).

The microclimate in pure eastern hemlock stands is substantially cooler than adjacent hardwood stands because of hemlock’s dense crowns. They grow in a wide variety of soil textures but most commonly in soils with impeded drainage (Tubbs, 1995). Hemlocks can persist longer than their more competitors and do not relinquish space easily.

Hemlock forests dominated by large trees on sheltered sites represent some of the oldest and more mature stands across southern New England (Foster and O’Keefe, 2000).

There are ~930,000 ha of hemlock-dominated stands (Tsuga spp.) in North America.

Associations may occur with other conifers and broadleaf tree species, such as the sugar maple (Acer sacharum Marsh.), American beech (Fagus grandifolia Ehrh.), yellow birch

(Betula allegheniensis Britton), and American basswood (Tilia americana L.). They grow on a diverse range of sites but generally are associated with riparian environments.

Hemlock’s natural range extends from northeast Minnesota across Wisconsin, northern

Michigan, south-central Ontario to the extreme southern Quebec, and through New

Brunswick and Nova Scotia. It is also found throughout New England, New York,

Pennsylvania, and as far south as northwestern Alabama (Godman and Lancaster, 1990.

Eastern hemlock is second only to sugar maple in abundance within the northern forest,

(Curtis, 1959). Old stands are divided between New England, with 54% of the hemlock area, the Mid-Atlantic states with 44%, and southern Appalachian states with 2%

(McWilliams and Schmidt, 2000). The total net volume of hemlock species (Tsuga spp.)

in the US is ~238 million cubic meters, representing 2% of the forest inventory within its

natural range. About 96 million cubic meters of live hemlock species are found within the

mixed maple-beech-birch forests. Around 60% of the harvested hemlock wood is used for pulp and paper production; the remaining is used for lumber (McWilliams and

Schmidt, 2000).

A dramatic decrease in hemlock forests has been quantified and mapped from

1984 to 1994 using remote sensing techniques (Royle and Lathrop, 2002). Half of the range of hemlock along the eastern seaboard were infested with Adelges tsugae (Knauer et al., 2002). Tree mortality occurs five to six years after a heavy infestation (Mayer et al., 2002). Extended research on the impact of this pest on eastern hemlock has been reported (Bonneau et al., 1999; McClure and Cheah, 1999; Colbert et al., 2002; Orwig,

2002). The subsequent impact of hemlock mortality due to this pest in the ecosystem has been also documented (Orwig et al., 2002; Kimple and Schuster, 2002; Williams et al.,

2002; Yorks, 2002). The most preoccupying characteristic related to the A. tsugae is its chronic nature. In contrast with other pest insects, A. tsugae persists, with no decline in the population after an outbreak (Souto and Shields, 2000).

The adelgid was first reported in the US in the Pacific Northwest in the 1920s. It apparently came from Japan where it is native, and was not a problem (Souto and

Shields, 2000). It was found in Virginia in the early 1950s and spread into New England

in 1985. It is now widely established in practically all eastern coast states (Fig. 1.1),

spreading at a rate

of 20-30 km/yr,

weakening and

destroying hemlock

stands in its wake

(Orwig and Foster,

1998).

In spite of the

Figure 1. Distribution of Hemlock Woolly Adelgyd, Adelges tsugae, in the FigureFigure 1.1.1.1.DistributionDistribution of hemlock of woolly Hemlock adelgid, Adelges Woolly tsugae , Adelgid, obvious importance easternin the eastern US (USDA US (USDA Forest Forest Service, Service, 2004). 2004). Adelges tsugae, in the eastern US (USDA Forest Service, 2004). of A. tsugae, the

elongate hemlock scale, Fiorinia externa (Homoptera: Coccoidea: Diaspididae), is also

an obvious stressor factor responsible for the decline of hemlock which has been, so far,

relegated to a secondary role as to the factors responsible for the current health situation

of hemlock forest in the northeastern US.

The elongate hemlock scale is an armored scale. Armored scales are the most

highly specialized plant parasites within the Coccoidea and usually attack perennial trees.

They are also pests of fruit, nuts, shrubs and ornamental plants, including greenhouse and

indoor plants. The worldwide production costs attributed to scales are estimated at 5

billion dollars annually, 500 million within the US (Kosztarab, 1996). Few countries in

the world today are exempt of diaspidids pest problems, which represent a serious

menace to the citrus industry in most countries where this crop is cultivated (Jeppson and

Carman, 1960). Their ability to successfully invade new territories has made them a

threat in many parts of the world (Morrison and Renk, 1957; Morrison and Morrison,

1965; Russel et al., 1974; Kosztarab and Kosztarab, 1988). Extensive research covering

all aspects of armored scales has been previously compiled (Helle, 1990 a, b).

F. externa is native to Japan (Takagi, 1963) and was accidentally introduced into

the US in the 19th century. In Japan it is commonly found on Tsuga diversifolia Masters

and T. sieboldii Carrière, only seldom causing damage (McClure, 1997). The Fiorinia

scale was first collected in the US by F. N. Meyer on Tsuga sp. from Queens, Long

Island, NY in 1908, and recorded as F. japonica Kuwana (Sasscer, 1912). It was later

described by Ferris (1942) as a new species from material collected by H. S. McConnell

in the area of Towson, north of Baltimore, MD, where the diaspidids were a local

problem. The scale was also present in Ohio (1929), Connecticut (1929), Pennsylvania

(1946), New Jersey (1950) and Washington, D.C. (1952) according to the United States

National Museum records (Garrett and Langford, 1962b).

The greatest

abundance of F. externa

occurs within a 300 km

radius of New York City

(Danoff-Burg and Bird,

2002), spreading from

southern New England

to western Ohio and Figure 1.2. Distribution of Elongate Hemlock Scale, Fiorinia externa, in the Fig. 1.2.FigureDistribution 1.2. Distribution of elongate of Elongate hemlock Hemlock scale, Scale,Fiorinia Fiorinia externa externa, in the, in the Tennessee (Fig. 1.2). Easterneastern US (Johnson US (Johnson and andLyon, Lyon, 1988; 1988; Hoover, Hoover, 2003; 2003; Lambdin Lambdin et et al.,al., 2005).20052005).).

The primary hosts are T. canadensis, T. caroliniana, T. diversifolia, Abies spp. and Picea spp., although it also feeds on Cedrus spp., Pseudotsuga menziesii, Pinus spp., and Taxus spp. (Hoover, 2003). The main host is eastern hemlock (McClure, 1988) but it can mature and reproduce on 57 different species among the native and exotic conifers

(McClure and Fergione, 1977), although infested hemlocks are always near. Outbreaks of this pest often intensify following infestation of A. tsugae, drought or other stress factors.

It may also co-exist with the circular scale [Nuculapsis tsugae (Marlatt)], another exotic species from Japan, however F. externa is a superior competitor due to the nutritional advantage gained by colonization of the tree earlier than its competitor hence monopolizing younger nitrogen-rich needles (McClure, 1980a; Reitz and Trumble, 2002).

1.2. The Fiorinia externa (Hemiptera: Coccoidea: Diaspididae) armored scale

Armored scales (Diaspididae) are characterized by a dorsal protective cover or shield, called the scale. It is composed of loose fibers secreted by pygidial glands, a fluid material discharged from the anus (thought to bind the fibers together) and two larval exuviae (one in the males), which are incorporated into the scale at each molt (Dickson,

1951; Disselkamp, 1954; Stoetzel, 1976). In the males the cover is a waxy fragile material and in the females it is hard and impermeable (Fig. 1.3).

In addition to the dorsal scale, most diaspidids produce a membranous ventral scale composed of secretions from the ventral pygidial glands, plus incorporated ventral exuviae residuals (Beardsley and Gonzalez, 1975; Kosztarab and Kozár, 1988). The shape of the scale, its color and the position of the exuviae are relatively constant in a given species, and these characteristics are valuable for identification purposes (Gill,

1997). In F. externa, the waxy covers can be observed on the lower surface of needles and on new cones (Johnson & Lyon, 1988; Hoover, 2003).

Figure 3. F. externa adult female carrying eggs under translucent exuvia (A) and F. Figure 1.3. F. externa adult female carrying eggs under translucent exuvia (A) and F.externa externaadultadult male male with with profuse profuse filamentous filamentous waxy strandsstrands(B). B). (B).

A distinct sexual dimorphism is characteristic in diaspidids. The F. externa female is elongate, with margins almost parallel and tapering at the apex. The length of the adult varies depending upon the stage of development, from 1.4 mm in a newly- molted individual to 0.6 mm in a female that has completed oviposition (Wallner, 1965).

The scale cover of the female is composed almost entirely of the sclerotized and elongate second exuvia (Ferris, 1942). The 2nd instar becomes heavily sclerotized, and the adult

female within remains membranous. The female is found within the exuvia of the 2nd instar known as the pupillarial form (Sasscer, 1912; Davidson & McComb, 1958;

Kosztarab, 1963). The adult uses the 2nd instar to protect the eggs in the same way that

non-pupillarial forms use the scale covering (William and Watson, 1988). The female is

more elongate than the male. The newly-molted adult female is pale yellow and gradually

darkens from pale to dark brown whereas the male is whiter and narrower in shape

(Duda, 1957; Wallner, 1965). The exuviae is translucent (Ferris, 1942). The old rostralis

is left in the plant tissue. The scale cover of females is yellow-brown or tannish and is

2 mm long; in males it is slightly smaller (1 mm) and whiter than the females (Wallner,

1965). The covering will not drop off even after death, though it will erode with time.

“The cover shows a specific peculiarity in that the first exuvia is exceedingly thin and transparent and appears to be detached from the second, to which it is connected only by a thin film of wax” (Ferris, 1942). Eggs (0.25 - 0.37 mm long) are laid under the 3rd instar exuvia. As egg laying progresses, the body of the female shrivels (Wallner, 1965).

Nymphs (crawlers) are flattened dorso-ventrally, 0.33 mm in length and are pale yellow-orange (Fig. 1.4).

On the dorsal surface of the 2nd instar exuvia there are

two long-necked glands that secrete wax (Wallner,

1965). Between the antennae (five segments, the distal

being as long as the other combined), two other glands Figure 1. 4. F. externa crawler. are present that secrete large amounts of filamentous

wax, commonly called silk (Stoetzel, 1976). A glutinose substance is then secreted by the

Malphigian tubules and discharged from the anus. This glues the filaments into a solid

mass that hardens and forms an elongate cap over and around the insect’s body

(Baranyovits, 1953; Stoetzel, 1976) which turns dark with age (Fig. 1.5). When these secretions are abundant, F. externa may be misdiagnosed as A. tsugae or spider mites

(Hoover, 2003). Three pairs of legs are present. The rostralis is located between the first

pair of legs (Wallner, 1965).

Crawlers are able to move a few meters, but once settled the 2nd instar starts

feeding, becomes robust with a lemon yellow coloration and loses its legs. Females

remain anchored for life (Baranyovits, 1953). The rostralis is composed of coiled stylets

that are three or four times the length of the body (Johnson and Lyon, 1988). Males and females are not morphologically distinguishable up to the 2nd instar stage, which is the

last feeding instar for males (Wallner, 1965).

“The third instar male scale is

elongated (0.49 mm), white, with

a terminal exuvia. The prepupal

male is orange-yellow in color

with 0.8 mm long. The antennae,

wings and legs appear as everted

sacs. The pupal male (0.85 mm) Figure 1.5. Settled F. externa nymph secreting glutinose substance from Malphigian tubules and young adult female with full developed is dark orange in color and the fresh scale cover. appendages are hyaline. The fully developed male, prior to emergence, is contained under a scale cover composed of the first instar exuvia and a filamentous, white covering. It has formed new appendages: two pair of wings, the second one reduced to haltere-like structures, antennae and three pairs of legs and a conical head” (Wallner, 1965).

1.2.1. Life-cycle

F. externa overwinters as a fertilized female or in the egg stage. In early spring

females start to deposit eggs beneath their waxy cover and may continue to lay eggs

throughout the summer, only interrupted by cold weather (Johnson and Lyon, 1988;

Hoover, 2003). One female can produce up to 20 eggs over the course of its life, though

the average at any one time is six (Davidson and McComb, 1958; Stimmel, 1980).

The female contracts and occupies the anterior half of the last instar exuvia, providing space for the eggs (Wallner, 1965). Egg production is greatest from late April through

May (Stimmel, 1980).

In 3-4 wk eggs hatch (early May) into nymphs (crawlers) that exit through an opening at the posterior end of the scale (Malinoski and Davidson, 2002; McClure,

2002b). They are very active and migrate preferentially to new needles (Davidson and

McComb, 1958; Stimmel, 1980). Living crawlers have been observed at a distance of 105 m from infested trees (McClure, 1977b). Within 1-2 days after hatching, crawlers settle in the underside of a needle and insert their stylets in the needle, which penetrates between the epidermal cells or through a stomatal opening where the insect feeds on the mesophyl cells (Wallner, 1978; Hoover, 2003). The first instar nymph starts to secrete the waxy cover around it as it grows. Only the head region of the insect remains uncovered

(Beardsley and Gonzalez, 1975).

Two types of waxy covering are produced: a frosty appearing layer from the posterior end which eventually covers the 1st and 2nd instars, and also a mass of trailing threads from two glands located between the antennae (Wallner, 1965). This latter secretion gives the tree the white-washed, mealy appearance. After 2-4 wk the insects’ body becomes flattened and the dorsal skin hardens and thickens to form the first scale cover (Wallner, 1965). They then molt into 2nd instars, and another 4 wk are required to reach maturity (Johnson and Lyon, 1988). Adults from this generation appear to reach maturity from mid to late July (Stimmel, 1980). The second stage male molts into a non- feeding prepupa and spins a cocoon, within which it pupates. When mature, males emerge as tiny winged insects, mate with the females and then die without feeding.

Mated females start to lay second-generation eggs (if present) six wk after mating

(Hoover, 2003). Eggs produced later (June or July) hatch and mature in a shorter period

of time (Stimmel, 1980).

F. externa completes two generations each year in the southern and Mid-Atlantic

States, but usually only one in the Northeast (McClure, 2002b). The climate of the East

has moderated in some areas, which may have allowed F. externa to become bivoltine in

this region as well (McClure, 2002a). A summer and an incomplete autumn generation

has been reported in Connecticut (McClure, 1977b, c) and New York (Wallner, 1964),

two generations in Massachusetts (Bray, 1958) and Connecticut (Talerico et al., 1967)

have been reported and multiple generations in Maryland have been also documented

(Davidson and McComb, 1958; Garret and Langford, 1969a). Overlapping stages can

indicate either unsynchronized single generations or multiple overlapping generations

(Stimmel, 1980). When a second generation occurs, individuals that develop from the eggs mature and a second outbreak of crawler occurs in September (Malinoski and

Davidson, 2002). This new generation overwinters as fertilized females. Adult females may live up to one year (Johnson and Lyon, 1988).

1.2.2. Damage

Leaf chlorosis and other localized toxic effects are commonly associated with armored scale infestations (Beardsley and Gonzalez, 1975). F. externa feeds on the needles of their hosts by removing fluids from the mesophyll cells through piercing- sucking mouthparts (Wallner, 1965). The younger needles of the lower crown are highly preferred as feeding sites, but the scales can be found on needles of every age and throughout the crown (McClure and Fergione, 1977). When populations are high, F.

externa causes a chlorotic appearance of the upper needle surface, which is related to the

destruction of mesophyll cells. Premature needle drop follows. In an advanced stage of

infestation there will be thinning foliage. Dieback of major limbs, progressing from the

bottom of the tree upwards usually occurs after F. externa densities reach 10 insects per

needle (McClure, 2002b). The excessive loss of plant fluids will debilitate the tree and

favor other pests such as the hemlock borer (Melanophila fulvoguttata Harris) or root rot

(Armillaria spp.) and are readily broken or thrown by wind (Hoover, 2003). Death of the

tree may occur in ~10 yr if populations are not controlled (McClure, 1980a).

1.2.3. Management

The management of accidentally introduced species is a difficult task. They

frequently represent the most serious insect pests as they lack natural enemies capable of

regulating their populations and maintaining them below an economic threshold. They

rapidly spread throughout a defenseless host. The usual approach to the problem is to use

insecticides or introduced natural enemies from the native geographic origin of the pest.

Both of these options have limitations, challenges and potential negative environmental

implications. Alternatives, such as extensive logging to remove infested trees, are

impractical in many hemlock stands, as access is difficult and not cost effective. Systemic

insecticides, such as Merit® (Imidacloprid), which are used to treat individual trees to

manage A. tsugae, are not effective in managing armored scale insects (Hoover, 2003).

Tree health is directly correlated to the potential impact of a pest population, particularly in piercing and sucking insects such as F. externa. Hemlocks have shallow roots and thus are susceptible to drought stress (Fowells, 1965). Hence, hemlocks located within an infested area should be maintained in good health to discourage pest problems.

Applications of nitrogen fertilizer and broad-spectrum insecticides can aggravate the pest

problem. Nitrogen enhances the survival, development rate and fecundity of scales,

resulting in higher densities on fertilized trees (McClure, 1980b). Residues from

insecticidal and fungicidal sprays and dusts in citrus foliage have been correlated with an

increase of purple scale Lepidosaphes beckii (Newm.), and the Florida red scale,

Chrysomphalus aonidum (L.) (Osburn and Spencer, 1938). Inert residues on leaves and

fruit enabled the young crawlers to become established by protecting them from direct

sunlight and heat, affording them a foothold among the residual particles.

In addition, an inadequate application of pesticides can cause resurgence of scale

populations by eliminating natural enemies. The elimination of the primary parasitoid of

the scale and three species of common predators from the lower crowns of partially

treated trees after insecticide treatment has been recorded (McClure, 1977a). Attempts to

find a practical solution for the management of hemlock scale have been tested for some

decades with variable results.

1.2.3.1. Chemical control

Excellent control of F. externa crawlers was obtained with a malathion-DDT

mixture and malathion sprays, (Davidson and McComb, 1958; Bray, 1958; Garrett and

Langford, 1969b). However, the number of applications needed throughout the intermittent emergence period of crawlers made this control measure impractical.

Control for up to 3 yr was reported when Dimethoate was applied as a foliage drench to the crowns of infested hemlocks (Wallner, 1962, 1965; Garrett and Langford, 1969b;

McClure, 1977d). However, even if effective for ornamental trees, there are many negative impacts associated using insecticides, and because hemlocks are commonly

16 found near bodies of water, insecticides are not suitable for many environmental conditions. Complete coverage of a mature tree is difficult even spraying from underneath to reach the scales (Stimmel, 1980). Rapid resurgence of the scale after insecticide application commonly occurs (McClure and Fergione, 1977). Evidence has shown a density-dependent feedback affecting the scale abundance by which dense scale populations significantly limit the success of individuals of the following generations and high mortality of crawlers following pesticide spraying allows a rapid resurgence of the scale population. Observations also confirmed that scales which survived the treatment had significantly higher fecundity than untreated controls, probably because of improved host quality followed by a reduced herbivore pressure (McClure and Fergione, 1977;

McClure, 1978a, 1979a).

1.2.3.2. Natural enemies: parasites and predators

The most important natural enemy of the Fiorinia scale is the aphelinid parasitoid, Encarsia citrina Craw [=Aspidiotiphagus citrinus Howard] (Hymenoptera:

Aphelinidae) (Fig. 1.6). High parasitism rates in the native area of the scale, Japan, have been observed, with mortality rates ranging from 90.2 - 94.2% (McClure, 1986). This parasitoid maintains the scale at innocuous densities in Japan, however, similar results do not occur in the northeastern US though the parasitoid has been established for several decades and parasitism rates have intensified as scale populations have increased

(McClure, 1977c). Lack of synchrony between the life cycles of the parasitoid and the host is believed to be the reason for the failure (McClure, 1978b).

FigureFigure 1.6 .1.6. Parasitized– Parasitized scalescale with with Encarsia Encarsiasp. (A)sp. and(A) single and singleEncarsia Encarsiasp. parasitoidsp. parasitoid(B). (B).

Moderate numbers of the native lady beetle, Chilocorus stigma Say, (Coleoptera:

Coccinellidae), have been recorded in infested areas of F. externa (Wallner, 1965). The

adults and larvae attacked all stages of the scale. However, predation was not insufficient

to control the scale population (Wallner, 1965). Recently, Cybocephalus nipponicus

(Coleoptera: Nitidulidae), a native species of China and Korea, reared by the New Jersey

Dept. of Agric. to control the Euonymus scale [Unaspis euonymi (Comstock)], has been

introduced in Pennsylvania to test its potential for the control of F. externa (Blumenthal,

2003). The movement of this predator from the Euonymus scale to feed in other scales,

including the Fiorinia scale had been observed. Releases and evaluations in Pennsylvania

hemlock forests indicate that the beetle has the potential to be effective, over time, in

controlling F. externa (Blumenthal, 2003).

1.2.3.3. Entomopathogenic fungi

The usefulness of fungi attacking scale insects has been exploited and studied

extensively (Gossard, 1903; Rolfs, 1908; Berger, 1919, 1921; Watson and Berger, 1937;

Fawcet, 1948; McCoy, 1985). Observations describing the occurrence of

entomopathogenic fungi on scales have been reported since the end of the 19th century from all continents (Table 1.1). In the 20th century, 37 parasitic and semi-parasitic fungi

18 were reported on scale insects (Fawcett, 1948). The presence of entomopathogenic fungi specifically isolated from Fiorinia spp. also has been observed since the beginning of the

20th century (Table 1.2).

In 2001, an entomopathogenic fungus attacking F.

externa and N. tsugae was reported (October 2001)

in the hemlock forest at the Mianus River Gorge

Preserve in Bedford, NY where an epizootic within

the scale population was observed (McClure, 2002).

FigureFigure 1.7.7 – F.F. externa externa imagoeimagoe presenting presenting Large sclerotia masses where found concealing in sclerotia masses. sclerotia masses. many cases the body of adult scales (Fig. 1.7).

No pathogens parasitizing F. externa had been previously reported. A group of entomopathogenic, phytopathogenic and endophytic fungi associated with F. externa, were isolated at the Entomology Research Laboratory, ERL (Marcelino et al., submitted).

However, one fungus was the most commonly retrieved pathogen in the 36 sites from

New York, Pennsylvania, Connecticut and New Jersey states, where collections of infected F. externa adults were made. This fungus was successfully inoculated in F. externa. Subsequent recovered of the fungus from infected dead scales revealed a potential use of this fungus for biological control.

Colonies of this fungus present a remarkable morphological plasticity, growing in radial non-uniform patterns and producing different pigmentation ranging from whitish- pink to intense black. Black hemispherical sclerotia, approximately 2 mm wide have been observed in the field. SDYA 1/4 media (Sabouraud's dextrose yeast agar 1/4 strength), with low concentration of carbohydrates, causes formation of the same sclerotious bodies

found in the field, after four days of incubation. Conidial spores have an elliptical shape

(6.51-6.73 × 3.26-3.35 µm) initially with round edges in young cultures and

progressively acute as the culture reaches full growth (Gouli et al., 2004). The genus was

preliminary identified by scientists at the Entomology Research Laboratory, UVM (S.

Gouli and V. Gouli, pers. comm. 2003) as Hypocrella sp. and the species was verified as

the anamorph Aschersonia marginata Ellis and Ever (Ascomycota: Hypocreales:

Clavicipitaceae) by Dr. Zengzhi Li (Head of Entomogenous Fungi Branch of the

Mycological Society of China). Doubts concerning the identification remained after contacts with worldwide renown taxonomists (Table 1.3 for identifications), since

Aschersonia spp. mainly grows in tropical or sub-tropical climates (see Appendix 1 for genus description), very different from the northeastern US climate. Molecular analysis allowed an accurate identification of the genus as Colletotrichum.

The genus Colletotrichum Corda (Ascomycetes: Phyllacorales: Phyllachoraceae) described by Sturm (1831) comprises one of the most extensive fungal groups of phytopathogenic species in the world, being a direct cause of plant anthracnose and damping-off, blight and spot disease in infected plant tissues, fruits, stems, roots and flower petals (Latunde-Dada, 2001). Entomopathogenic isolates of Colletotrichum spp. have only been reported to occur on Orthezia praelonga Douglas (Hemiptera:

Ortheziidae), a major scale pest of Citrus spp. in Brazil (Robbs et al., 1991).

Colletotrichum gloeosporioides causes an intense epizootic in populations of this insect

(Cesnik and Oliveira, 1993) confirming an early Brazilian report (Batista and Bezerra,

1965).

The ubiquitous presence of this unreported entomopathogenic strain of

Colletotrichum in the epizootic areas led us to focus our attention and research in the

biological processes involved in the interactions between the insect pest (F. externa) and

fungal pathogen (Colletotrichum sp.) and to determine the origin and biocontrol potential

of this entomopathogenic fungus.

1.3. Entomopathogenic fungi as biological control agents

Pathogens have long been known to play a major role in the population dynamics

of many important forest pests. Among the causal agents of diseases in insects, such as

protozoans, bacteria, viruses, rickettsia and nematodes, the entomogenous fungi

(entomophatogenic and entomoparasitic fungi), obligate or facultative parasites of insects, play a relevant role. Because of their conspicuous macroscopic growth on the

surface of their hosts, fungi were the first microorganisms found to cause disease in

insects (Tanada and Kaya, 1993). Economically, the use of entomopathogenic fungi

offers a unique alternative to a chemical control approach of an insect pest. Under ideal

conditions an area of 1 ha can be treated with fungal formulations at a cost of $20

(Wraight et al., 2001). In addition, nontarget safety testing increases the registration cost of a chemical pesticide to an estimated $20-40 million (Marrone and MacIntosh 1993), contrasting with the approximately $2 million for a biological control agent (Rodgers,

1993). Successful biocontrol has been extensively reported (Campbell et al., 1985;

Fransen, 1987; Lewis et al., 1996; Lacey et al., 1996; Charnley, 1997; Keller et al., 1997;

Inglis et al., 2001; Meekes et al., 2002) as well as commercial myco-insecticide use (see

product list in Baron, 2002; IR-4, 2005). The most outstanding achieved control, up to

21 now, was the introduction and establishment of the entomopathogen Entomophaga maimaiga Humber, Shimazu & Soper, which has caused a massive epidemic in northeastern US gypsy moth (Lepidoptera: Lymantriidae) populations. This pest has been maintained below the economical threshold for several years due largely to this fungus

(Hajek et al., 1996; Dwyer et al., 1998). There are at present 74,000 species of fungi identified worldwide (Hawksworth, 2001), a portion of which are insect-specific

(Rossman, 1994). The number of fungi associated with insects is so vast and their role so relevant, that their inclusion in biodiversity projects is of primary importance (Benjamin et al., 2004). Exploitation of pathogenic organisms, under suitable environmental conditions, may be an important tool to maintain an insect’s population under an economic threshold. If the parasitic pathogen subsequently establishes and successfully maintains its viability in the environment, it can become a practical pest management option.

Records of entomopathogenic fungi from the Oligocene-Miocene boundary (~ 25 million yr old) have been reported, indicating fungi pre-date human existence (Poinar and

Thomas 1982, 1984). In the late 19th century, research directed towards the taxonomy of entomopathogenic fungi increased. Petch (1870-1948) in Asia, and Mains (1890-1968) in

America, led the field of fungal taxonomy (Smith, 1969). Intensive research focusing on the use of fungi for the control of agricultural pests began in the 20th century (Samson et al., 1988). This period was followed by a decline phase in the mid-1950s. Synthetic organic pesticides, epitomized by DDT, became the dominant control technique after

1945, especially for the control of forest insect pests (Perkins, 1988). Synthetic insecticides have remained the primary control method but increased attention directed

towards biological control methods began in the 1980s (Ignoffo and Anderson, 1979;

Burges, 1981; Hall and Papierok, 1982; Jaques, 1983; Rombach and Gillespie, 1988),

stimulated by the general awareness of insect resistance to chemical pesticides and the

environmental hazards of chemical applications. The potential dangers of pesticides were

reported by governmental agencies, including the National Research Council of the

National Academy of Sciences, and by various congressional committees (Knipling,

1979). Many popular books on the subject were written, including Rachel Carson’s

‘Silent Spring’ (1987).

Examples of successful biological control with bacteria and fungi have been

reported in North America (Burges and Hall, 1982) and England (Wilding, 1983). At

present, the commercialization of mycoinsecticides is no longer a pioneering field but a

common resource. The forestry market was one of the first in which a microbial agent

(Bacillus thurigiensis subsp. kurstaki) replaced broad-spectrum chemical insecticides

(Frankenhuyzen et al., 2000). Sales of the bacterium Bacillus thuringiensis for insect

control represented over 90% of the international biocontrol market in the 1990s

(Rodgers, 1993).

Of the ~ 700 species of entomopathogenic fungi distributed in 100 genera, a few

are in use or under development for insect control (Lisansky and Hall, 1983; Roberts and

Wraight, 1986, McCoy et al., 1988) in agriculture, forestry or medical fields. Genera

containing entomopathogenic species were compiled in the ‘Atlas of Entomopathogenic

Fungi’ (Samson et al., 1988) and by McCoy et al. (1988). Later Humber (1989, 1992,

1997 and 1998) and Mueller et al. (2004) compiled the most characteristic conditions in

which a fungus might occur to cause infection in insects, which is spore-bearing states

(Fig. 1.8).

Entomopathogenic fungi

Deuteromycota Deuteromycota (Fungi imperfecti) (Fungi imperfecti) Zycomycota Mucorales

-Aschersonia Mont -Cordyceps Fries -Myiophagus Thaxter -Beauveria Vuillemin -Hypocrella Saccardo ex Sparrow -Fusarium Link: Fries -Lagenidium Schenk -Coelomomyces Keilin -Gibellula Cavara emend. Humber and Rombach -Hirsutella Patouillard -Hymenostisbe Petch emend Samson and Mucorales Enthomophthorales Evans -Metahrizium Sorokin -Nomurea Maublanc -Sporondiniella -Batko Humber -Paecilomyces Bainier Boediin -Conidiobolus Brefeld -Pseudogibellula -Entomophaga Batko Samson and Evans -Entomophthora -Septobasidium Fresenius Patouillard -Erynia Nowakowski -Sorosporella Sorokin -Furia (Batko) Humber -Tolypocladium Gams -Massospora Peck -Verticilium Nees per -Neozygites Witlaczil Link -Pandora Humber

Adapted from:

Samson et al. (1988); McCoy et al. (1988); Humber (1989, 1992, 1997 and 1998); Mueller et al. (2004).

Figure 1.8. Taxonomic classification of entomopathogenic fungi

The phylum Deuteromycota contains the largest number of entomopathogenic fungi, including genera with a wide geographic distribution (Beauveria spp., Metarhizium spp.), comprising primarily mycopathogens of soil insects, i.e. Beauveria bassiana

(Bals.) Vuill. and Metarhizium anisopliae (Metsch) Sor. (Daoust and Roberts, 1982,

1983; Fargues and Robert, 1985; Keller and Zimmerman, 1989).

Species with a main tropical or subtropical distribution (Gibellula spp. and

Hymenostilbe spp.) and genera presenting host group specificity, such as Nomuraea rileyi

(F.) Samson to Lepidoptera, or Aschersonia sp. to scales and whiteflies are also present in

the Deuteromycota, (Zimmermann, 1986; Humber, 2000). The Basidiomycota

(Septobasidium sp. Pat.) are absent from the list since evidence indicates they essentially

live symbiotically with the insect (Samson et al., 1988). Septobasidium sp. constitutes a distinct evolutionary shift from the common entomopathogenic pattern of activity of fungi where death overcomes quickly after host infection (Humber, 1984). However, pathogenic representatives of Septobasidium sp. are known (Evans, 1989).

For a thorough understanding of the principles of insect pathology, a detailed knowledge of the structure and biology of the infective agent is imperative. Unlike other insect pathogens, entomogenous fungi initiate infection by attachment and germination of the spore which is able to penetrate the cuticle of the host insect. Infection from ingested spores is a rare occurrence (Roberts and Humber, 1984). Entomopathogenic fungi have spores adapted to attach to an insect host, but the physical and chemical characteristics of spore and cuticular surfaces responsible for attachment are unknown (Roberts and

Humber, 1981). The adhesion of the fungal spore to the insect cuticle is a critical phase in the infection process. Hypovirulence can result from a deficient attachment of spores.

Many aspects are involved in the adhesion process such as the physiological state of the host, chemical nature of the cuticle surface, microclimatic conditions of host cuticle and host specificity (Boucias and Pendland, 1983; Farges, 1984).

After adhesion, the penetration of the insects’ cuticle (involving both toxin and enzyme production) occurs, either directly by germ tubes or by infection pegs which evaginate to form appressoria. After penetration into the insect’s body, a multiplication phase (hyphal bodies) takes place in the haemocoel (Roberts, 1981). After filling the host with mycelium, hyphae grow in the insect’s integument. Finally, production and dissemination of spores initiates on the external surface of the host (Poinar and Thomas,

1978). After death, the insect’s body becomes hard and mummified, containing the fungal stroma. Dead insects are generally attached to the subtract where they were infected (e.g. plants) by their appendages or proboscis, or by rhizoids or pseudocystidia produced by the fungus (Tana and Kaya, 1993).When an infection occurs naturally, without human intervention, such as the present case in study, and the entomopathogenic fungi rises to an epizootic level, control can be impressive. Unfortunately, this situation usually occurs after the economic threshold has been reached.

The current epizootic occurring naturally within population of F. externa in the northeastern hemlock forest of New York, Pennsylvania, Connecticut and New Jersey states, is an exceptional opportunity to study the etiology of the most commonly isolated microorganisms from infected scales within the geographic distribution of the epizootic,

Colletotrichum sp.

1.4. The genus Colletotrichum

Colletotrichum spp. are a direct cause of anthracnose in a vast number of plant species (Latunde-Dada, 2001). The common symptoms of disease are dark, sunken, lenticular necrotic lesions that occur in developing and mature tissues, usually containing

the acervuli of the fungus (Prusky et al., 2000; Latunde-Dada, 2001). Colletotrichum spp.

can also be saprophytic or secondary invaders of moribund tissue (Waller, 1992; Mills,

1994).

This genus is widely used as a model system for molecular and phytopathogenesis

studies comprising infection processes, resistance mechanisms, mycoherbicide use,

quiescence, biochemistry and molecular biology of host specificity (Prusky et al., 2000;

Rodriguez and Redman, 2000; Latunde-Dada, 2001). Colletotrichum and Phythophtora are the most studied plant pathogenic fungi in the world (Prusky et al., 2000). The revision by von Arx (1957) was a relevant milestone in the taxonomy of the genus, reducing the existent 750 species to 11 taxa based on morphological characteristics. The use of DNA techniques allowed a more accurate examination of the different species and taxa considered by von Arx (1957). Host specialized forms were identified as distinct species by Sutton (1992) who reported a total of 39 species in this genus. At present there is a lack of consensus in the scientific community regarding the number of species in the genus Colletotrichum (Cannon et al., 2000). It is currently undergoing intensive revision and clarification with the use of molecular taxonomic techniques. Species complexes and subspecific groups recently have been proposed (Latunde-Dada, 2001).

1.4.1. Taxonomic considerations

Genetic and mating type studies indicate that the population structure and dynamics of Colletotrichum spp. is complex (Chacko et al., 1989, 1990; Cisar et al.,

1994, 1996; Roca et al., 2004a, 2004b). Sexual compatibility and outcrossing between isolates of the same species and between species with different host specificities are the likely cause of the high degree of variability observed (Baxter et al., 1985; Cisar et al.,

1994). Genotypic diversity is enhanced mainly by outcrossing rather than inbreeding

(Frank, 1992).

Morphologically-similar races of a Colletotrichum sp. pathogen are characterized

in the basis of physiological parameters such as adaptation to different hosts (see Ansari

et al., 2004). They are then separated into biotypes called formae speciales (Shoemaker,

1981; Cisar et al., 1994). These biotypes are considered to be genetically distinct,

although evidence of chromosome transfer suggests that genetic exchange between

biotypes can occur in the field (Masel, 1996). Several Colletotrichum spp. may be

associated with a single host and a single Colletotrichum sp. may infect multiple hosts

(Freeman et al., 1998). There is uncertainty whether Colletotrichum isolates from one

host can successfully colonize and infect another host or if they are host-specific

(Vaillancour and Hanau, 1992).

For example, Colletotrichum gloeosporioides f. sp. aeschynomene has been tested

for pathogenicity in 30 different plant species and 46 cultivars of economic relevance,

and also in wild plants in the Leguminosae, in order to assess the specificity, biological

control potential and commercialization potential of this strain against Aeschynomene

virginica (Templeton, 1984). The only susceptible species were the target weed A.

virginica, and a related weed, Aeschynomene indica (Daniel et al., 1973). The pathogen

was released commercially in 1982 as CollegoTM (Weidemann, 1991). However, TeBeest

(1988) tested this same Colletotrichum species in 77 plants comprising 43 genera in 10 families, reporting that five genera in the subfamily Papilionidae included susceptible species to the fungus, although susceptibility diverge, with only the target weed A.

virginica being killed. The relationship between plant species and the pathogen

specificity is unclear (Weidemann, 1991).

The complete host range of most Colletotrichum species is largely unknown

(Fagbola and Abang, 2004). Pathogens with a wide range of hosts, such as C.

gloeosporioides, can have special forms (formae speciales) with more limited host ranges

(Weidemann, 1991). Three major species aggregates of Colletotrichum are currently

recognized: C. gloeosporioides, C. acutatum and C. orbiculare. Further reassembling will

probably confirm two more groups, C. capsici and C. graminicola (Cannon et al., 2000).

The concept of species in the genus Colletotrichum is currently being debated

(Sutton, 1992). Morphological characteristics initially used to determine species within

the genus (i.e. shape and size of conidia, setae, and appressoria) are widely recognized as

inadequate (Sherriff et al., 1994; Sreenivasaprasad et al., 1996; Cannon et al., 2000). For

example, C. acutatum was considered a morphological variant and synonymous with C. gloeosporioides until the Simmonds revision of the genus in 1965 (Cannon et al., 2000).

In 1957 the genus was again revised and >600 synonyms of C. gloeosporioides were

found (von Arx, 1957). Many contained morphological and physiological differences.

The von Arx report was then revised and C. gloeosporioides was described comprising

seven formae speciales (Sutton, 1992). Wide morphological plasticity in culture

including pigmentation differences, growth rate and overlapping phenotypes have been

consistently reported (Baxter et al., 1983; Walker et al., 1991; Freeman and Rodriguez,

1995; Lardner et al., 1999; Prusky et al., 2000; Sanders and Korsten, 2003; Couto and

Menezes, 2004). Recently, vegetative compatibility groups (González and Sutton, 2004) and molecular analyses have been used to determine diversity within species (Freeman et

al., 1996; Freeman and Katan, 1997; Ford et al., 2004). Evolutionary relationships

between Colletotrichum lineages also have been studied, confirming the genetic diversity within this genus and the difficulties to clade species based on genetic parameters

(Crouch, 2006).

1.4.2. rDNA analyses to discern Colletotrichum spp.

In the last decade, development of molecular techniques, coupled with traditional methodologies have been used to clarify the phylogenic relationship within the genus

Colletotrichum (Mills et al., 1992; Johnston and Jones, 1997; Balardin et al., 1999;

Freeman et al., 2001a; Afanador-Kafuri et al., 2002; Ureña-Padilla et al., 2002; Talhinhas et al., 2002). As an example, sequence analysis of rDNA has enabled the identification of multiple genotypes of C. acutatum (Förster and Adaskaveg, 1998; Freeman et al., 1998) and also clonal sub-populations differing in colony appearance (pink vs. gray cultures), conidia morphology, virulence, temperature relationship for growth and genotype

(Adaskaveg and Hartin, 1997; Förster and Adaskaveg, 1998). Population diversity within this species in New Zealand from fruit rots, lupin and pine, led to the determination of C. acutatum as a species group (Colletotrichum acutatum sensu lato). Host specificity

(Lardner et al., 1999) and molecular analyses made it possible to distinguish four C. acutatum sensu stricto sub-groups (Johnston and Jones, 1997). Subsequently two additional distinct groups have been found (Guerber et al., 2003). A comprehensive molecular analysis of C. acutatum sensu Simmonds (one of the sub-groups within C. acutatum sensu stricto) has been conducted (Freeman et al., 2001). The phylogeny and systematics of 18 Colletotrichum spp. led to the conclusion that C. acutatum shows the

maximum intraspecific divergence in the ribosomal DNA internal transcribed spacer

(ITS) and C. capsici the greatest interspecific divergence (Sreenivasaprasad et al., 1996).

The cosmopolitan host range of this species is repeatedly assessed and its host range

expanded (Sreenivasaprasad and Talhinhas, 2005).

Analysis using Random Amplified Polymorphic DNA (RAPD) and Restriction

Fragment Length Polymorphism (RFLP) of the 28S, 18S, 5.8S ribosomal subunits

(coding and conserved) and ITS regions (non-coding and variable) have also been

conducted on fungal species identification processes (Bailey et al., 1996; Martinez-

Culebras et al., 2003; Fagbola and Abang, 2004). The ribosomal internal transcribed

spacers I and II (ITS1 and ITS2) have been widely used as a base for Colletotrichum

species differentiation (Freeman and Rodriguez, 1995; Freeman et al., 2000; Moriwaki et

al., 2002; Martinez-Culebras et al., 2003). The 18S and 28S regions are functional and

more conserved, although some parts (especially D1 and D2 domains) are known to be

variable (Cannon et al., 2000). Conserved gene regions can be used for phylogenetic

analysis at the genus level and above (Mindell and Honeycutt, 1990; Hills and Dixon,

1991).

The ITS regions play a role in forming the three-dimensional structure of

ribosomes, because these regions do not interact and display more variation. Thus, they are valuable tools for differentiation of taxa at and below the species level (Cannon et al.,

2000). In Colletotrichum spp. the ITS1 region, with 50.3% variable sites, has a greater intra-and inter-specific divergence than the ITS2 region with 12.4% variable sites

(Sreenivasaprasad et al., 1996). For differentiation within Colletotrichum spp. relatively low variation in ITS sequences may occlude intraspecies diversity. Therefore, the introns of the glutamine synthetase (GS) and the glyceraldehydes-3-phosphate dehydrogenase

(GPDH) genes have been used to identify genetic differences (Guerber et al., 2003).

These DNA regions are valuable phylogenetic tools to identify inter-and intra-specific diversity in Colletotrichum spp. (Liu and Correll 2000; Liu et al., 2001) and for marker gene expression and transformation studies with the teleomorph of Colletotrichum,

Glomerella (Templeton, 1992a). Molecular data are used to create phylogenetic trees that

approximate to the evolutionary history (Berbee and Taylor, 2001).

1.4.3. Hosts

1.4.3.1. Plants

The range of hosts within the genus Colletotrichum is wide, and reports of new

host plants are common. The genus shows a wider expression in warm moist

environments such as the humid and sub-humid tropical zones (Mills et al., 1992; Waller,

1992). Forty seven cultivated host species in South Africa have been reported to be hosts

of Colletotrichum spp. (Gorter, 1977 cit. in Baxter et al., 1983). In New Zealand 39 host

species from 23 plant families also have been reported (Simmonds, 1965). Xiao et al.

(2004) recovered Colletotrichum spp. from 23 cultivated and non-cultivated host plant

species in west-central Florida, US. In Japan, Moriwaki et al., (2002) isolated 25

Colletotrichum species from 123 plant species including Gramineae, Leguminosae and

Cucurbitaceae. An extensive record of hosts (grain legumes, pasture legumes, trees) was

made by Lenné (1992), assessing at least nine species of the fungus within 102 plant species.

Economically significant losses occur in cereals, grasses, legumes, woody plants, trees, vegetables and perennial crops worldwide as a result of Colletotrichum spp.

(Lardner et al., 1999). The complex of Colletotrichum spp. causing anthracnose disease is

the most serious threat to the strawberry industry (Fragaria x ananassa Duchesne).

Colletotrichum fragariae Brooks was first reported on strawberry in the US by Brooks

(1931). Later, C. acutatum Simmonds ex Simmonds (Simmonds, 1965; Smith and Black,

1986) and C. gloeosporioides (Penz.) Penz. & Sacc. in Penz. (teleomorph Glomerella cingulata [Stonem.] Spauld. & H. Schrenk.) were reported to cause anthracnose disease in strawberry, damaging roots, stolons, seeds, petioles, immature leaves, buds, fruits, flowers and causing crown rot in all cultivated strawberries worldwide (Kulshrestha et al., 1976; Gunnell and Gubler, 1992; Howard et al., 1992; Freeman and Katan, 1997;

Tanaka and Passos, 1998; Tanaka et al., 2001; Mertely and Legard, 2004; Horowitz et

al., 2004). Some of these diseases do not produce the dark sunken lesions characteristic

of anthracnose and a more general description of Colletotrichum strawberry diseases, is

proposed (Legard, 2000). Colletotrichum acutatum is a quarantine organism on

strawberry in the European Union (Parikka and Lemmetty, 2004). It has been a

quarantined organism in Canada until 1997 when this measure was deregulated

(Canadian Food Inspection Agency, 1997). The morphology and cultural characteristics,

infection and pathogenicity mechanisms of Colletotrichum spp. on strawberry have been

documented (Smith, 1990; Horowitz et al., 2004).

Colletotrichum gloeosporioides has been reported to cause losses of 35-70% in

sweet and hot peppers (Alexander and Marvel, 2001). Colletotrichum sublineolum causes

economical loses in sorghum (Khan and Hsiang, 2003) and Colletotrichum graminicola

(Cesati) Wilson causes anthracnose disease on several economically important crops

including maize (Nicholson, 1992; Bergstrom and Nicholson, 2000), annual blue grass

(Poa annua) and bent grass turfs (Agrotis spp.) on golf courses in the northern US,

Canada and Europe (Browning et al., 1999) and other turfgrasses (Khan and Hsiang,

2003).

Colletotrichum coccodes (Wallr.) S. J. Huges is an important pathogen in potato production causing anthracnose on tuber surfaces (Tsror et al., 1999; Lees and Hilton,

2003). It is also important in tomato production causing anthracnose in fruits and roots

(Byrne et al., 1997; Dillard and Cobb, 1998). Fifteen species of weeds growing between crop rotations have been found to serve as hosts for C. coccodes (Raid and Pennypacker,

1987).

A common characteristic of C. sublineolum, C. graminicola and C. coccodes is the development of black micro-sclerotia within infected tissue (Byrne et al., 1997).

Colletotrichum truncatum (Khan and Sinclair, 1992), C. cocoodes (Sanogo and

Pennypacker, 1997), C. acutatum and C. gloeosporioides (Dingley and Gilmour, 1977) produce sclerotia in culture. The latter two species produce, respectively, poorly developed sclerotia and abundant sclerotia.

Proteaceae plants, valuable commercially and sought after as cut flowers, are also hosts of Colletotrichum spp., causing seedling damping off, anthracnose, leaf and stem lesions (Lubbe et al., 2004). Colletotrichum trifolii is an important pathogen for alfalfa legume crops (Dickman et al., 1995). Colletotrichum truncatum causes yield losses of lentil in the northern US (Venette et al., 1994) and Canada (Ford et al., 2004). Lupin

(Lupinus spp.) crops valuable for their high seed protein content are also affected by anthracnose (Nirenberg et al., 2001; Talhinhas et al., 2002). Colletotrichum lindemunthianum (Sacc. & Magnus) Briosi & Cav., is a serious disease in beans

(Phaseolus vulgaris L.) with losses of 95% in susceptible cultivars (Guzman, 1979; Tu,

1992; Pereira et al., 1998; Melotto et al., 2000). Colletotrichum demantium is the causal agent of anthracnose in cowpea stems (Smith and Aveling, 1997).

Perennial crops are highly susceptible to Colletotrichum species. The most consistently reported hosts are the following: avocado (Freeman et al., 1996; Everett,

2003; Sanders and Korsten, 2003), banana (Couto and Menezes, 2004), cashew (Muniz et al., 1998), coccoa (Dodd et al., 1992), coffee (Sreenivasaprasad et al., 1993; Nachet and

Abreu, 2002), cotton (Bailey, 1996), kenaf and jute (Waller, 1992), mango (Afanador-

Kafuri et al., 2002; Sanders and Korsten, 2003), papaya (Waller, 1992), passiflora

(Afanador-Kafuri et al., 2002), pepper (Ivey et al., 2004), rubber (Fernando et al., 2000), sugarcane (Wallner, 1992), tamarillo (Afanador-Kafuri et al., 2002), tea (Waller, 1992) and tobacco (Waller, 1992). Colletotrichum gloeosporioides and C. acutatum are the most important pathogens. Fruit and nut producing trees with anthracnose caused by the fungus have been also reported: almond (Förster and Adaskaveg, 1999; Freeman et al.,

2000), peach (Adaskaveg and Hartin, 1997; Ureña-Padilla et al., 2002) and citrus

(Zulfiqar et al., 1996; Timmer and Brown, 2000; Moretto et al., 2001). Fruit-rots caused by Colletotrichum spp. were observed in apple, fig and pear (Johnston and Jones, 1997), olive (Martín and García-Figueres, 1999) and grapes (Talhinhas, 2002). Deciduous trees have been reported hosts. Canker and dieback of grafted Japanese maple (Acer palmatum) was reported in a nursery in Connecticut, US and the cause was an infection by C. acutatum (Smith, 1991b; Smith, 1993).

Conifer trees also have been reported as hosts. Colletotrichum acutatum was reported affecting four month old seedlings of western hemlock [Tsuga heterophylla

(Raf.) Sarg.], western red cedar (Thuja plicata Donn), and Sitka spruce [Picea sitchensis

(Bong.) Carr.] in British Columbia, Canada (Hopkins et al., 1985). A subsequent study in nurseries where infections were observed showed the disease was confined to 380 western hemlock seedlings (Hopkins et al., 1985). Colletotrichum acutatum f. sp. pinea was found to cause temporary stunting from which seedlings normally recovered within the first year (Canadian Food Inspection Agency, 1997). Surveys in outplanted sites with western hemlock seedlings on Vancouver Island revealed the presence of C. acutatum in

70,000 seedlings (Hopkins et al., 1985). Colletotrichum gloeosporioides was also reported to cause blight in ~250,000 greenhouse grown western hemlock seedlings (70 d old) near Vancouver, Canada (Lock et al., 1984). Disease susceptibility to C. acutatum was assessed in western hemlock and mountain hemlock, being 95% and 92%, respectively (Griffin et al., 1987). Seedlings of Pinus radiata D. Don were found infected

by C. acutatum f. sp. pinea Dingley & Gilmour causing terminal crook disease in New

Zealand (Dingley and Gilmour, 1977; Nair and Corbin, 1981; Nair et al., 1983).

Although initially it was suggested a spread of the pathogen from the co-cultivated tree lupin, Lupinus arboreus (used to provide nitrogen for young P. radiata plantations), pathogenicity tests assessed the host specificity, finding isolates collected from lupin were different from those collected from pine (Lardner et al., 1997).

1.4.3.2. Mammals and amphibians

Although rare, mycosis caused by C. gloeosporioides has been reported in humans, mainly opportunistic infection in previously open injuries (Guarro et al., 1998,

Cano et al., 2004). Keratitis has been reported to be caused by C. graminicola in humans, usually after eye injury (Ritterband et al., 1997) and a mycosis by C. acutatum in amphibians also has been observed (Manire et al., 2002).

1.4.3.3. Insects

Biological control of O. praelonga (Hemiptera: Coccoidea: Ortheziidae) with the

fungus C. gloeosporioides f. sp. ortheziidae has been used for the last 13 years in Brazil

(Robbs et al., 1990, 1991, 1993; Viegas et al., 1995; Cesnik and Ferraz, 2000). Mortality rates in this scale insect, following applications of suspensions of C. gloeosporioides f.

sp. ortheziidae, ranged from 42-82% after 35 d and 85-96% 70 d post-application (Cesnik et al., 1996). Assays were developed to evaluate the effect of the isolates on Citrus spp.

They were not pathogenic to field-grown Citrus spp. plants but were to these grown in

greenhouses, although the fungus was commonly recovered from treated plants in the

field (Teixeira et al., 2001). Peach, apple and pumpkin leaves were also evaluated and

determined to be susceptible to all isolates if they were previously injured (Teixeira et al.,

2004). No toxicity to Crustaceae (Jonsson and Genthner, 1997) or mice (Castro et al.,

1998) was found.

Scientists at Embrapa Meio Ambiente, Jaguariúna, Brazil, studied the control of

O. praelonga from 1992 to 2003 and reported they have found no other fungus with the same infectivity to O. praelonga as Colletotrichum during their intensive field testing

(Cesnik et al., 2005). The large scale production of the fungus has been limited by bureaucratic constrains due to the reports of phytopathogenic strains of this fungus

(Cesnik et al., 2005), however commercialization in underway (R. Cesnik, pers. comm.).

Studies on the biochemistry and phytopatogenicity of the entomopathogenic isolates from

Orthezia have shown a distinct divergence from the known phytopathogenic isolates

(Rosamiglia and Melo, 1996).

Benoit et al. (2004) recently reported isolation of C. acutatum from internal

mycoflora of the cave cricket, Hadenoecus cumberlandicus (Hemiptera:

Rhaphidophoridae), suggesting the pathogen might have been acquired during plant foraging. However, re-evaluation of the fungus showed no effect on insect survival

(Benoit, pers. comm.).

1.4.4. Biocontrol of weeds with Colletotrichum spp.

Weeds cause major problems in agriculture systems worldwide, reducing nutrient intake for vigorous crop growth and thus reducing the quality and market value in crops

(TeBeest et al., 1992). Five strains of Colletotrichum spp. have shown commercial potential and 19 strains have been researched as possible mycoherbicides (Morris, 1983;

Mortensen, 1988; Charudattan, 1991; Makowski and Mortensen, 1992; Templeton,

1992b). Biocontrol programs using Colletotrichum spp. as a mycoherbicide are being

developed in North America, Europe and Asia.

CollegoTM (Encore Technologies, US), a dry powder formulation with C.

gloeosporioides f. sp. Aeschynomene has been commercially available since 1982 for

control of Aeschynomene virginica (L.). This weed is responsible for crop losses in rice

(Oryze sativa L.) and soybeans (Glycine max L. Merr.) in Arkansas, Louisiana and

Missouri, US (Bowers, 1986; Templeton, 1992b). When compared to chemical

herbicides, CollegoTM was nearly as effective (Yang and TeBeest, 1993). From 1982 to

1988 farmers treated 8000 ha/yr of rice and soybeans (Smith, 1991a). Production was

stopped in 2001 due to market constrains (D. Goulet, pers. comm.). CollegoTM (active ingredient, Colletotrichum gloeosporioides f. sp. Aeschynomene) and DeVineTM (active ingredient, Phytophthora palmivora (Butler) Butler] (Charudattan, 1991) were the first

fungal herbicides introduced for commercial use in the US. Until 1996, only BioMal®

was registered for commercial use, applying another strain of Colletotrichum spp. as the

active ingredient (Hoagland, 1996).

BioMal® (Philom Bios Inc.) was registered in Canada in 1992, but has not been

commercially used (G. Hnatowich, pers. comm.). It has been tested for the control of

Malvaceae weeds in flax and lentils. The active ingredient of the dry formulation is C.

gloeosporioides f. sp. malvae (Makowski and Mortensen, 1992; Morin et al., 1996). The

management of Xanthium weeds, noxious to pastures and crops, with Colletotrichum spp.

has been investigated (McRae and Auld, 1988; Walker et al., 1991). Burr Anthracnose®

using C. orbiculare as the active ingredient was tested for control of spiny cockburr

(Xanthium spinosum L.) in Australia, but it is also not commercially available

(Templeton, 1992b).

Velvetleaf (Abutilon theophrasti Medik) is a major annual noxious weed in

soybeans and corn (Zea mays L.) in the US, Canada and Europe (Warwick and Black,

1988). Strains of Colletotrichum coccodes selectively attack these crops (Wymore et al.,

1988). C. coccodes was initially studied as an application combined with chemical herbicides. The mixture increased the spore viability of the fungus (Wymore and Watson,

1986; Wymore et al., 1987). Subsequent research focused on the effect of C. coccodes in the intra-and inter-specific competition potential between velvetleaf and soybean plants

(DiTommaso and Watson, 1995; DiTommaso et al., 1996). Colletotrichum coccodes, registered under the name Velgo® in Canada and the US is considered a successful

bioherbicide using a Colletotrichum sp. as active ingredient (Templeton, 1992b;

Mortensen, 1988; Dauch et al., 2002).

Colletotrichum truncatum Andrus & W. D. Moore has shown promising

bioherbicide potential against the weed Sesbania exaltata (Raf.) Cory (Silman et al.,

1993; Connick et al., 1996). Corn flour-formulations of C. truncatum micro-slerotia incorporated into potting soil enhanced disease in emerging sesbania seedlings (Jackson et al., 1996). Biological control of common St. Johnswort (Hypericum perforatum), an aggressive weed widely disseminated, can be enhanced using a host-specific strain of the fungus, C. gloeosporioides f. sp. hypericum (Penz.) Penz. and Sacc. (Morrison et al.,

1998). Colletotrichum gloeosporioides f. sp. malvae was submitted for review by Encore

Tech. to the EPA (US Environmental Protection Agency) for possible registration in the

US under the name Mallet W. P. but approval is still pending (G. Hnatowich, pers. comm.). Results with the use of C. gloeosporioides for control of lodgepole pine dwarf mistletoe (Arceuthobium americanum Nutt. ex Engelm.) are encouraging, and a worldwide patent is being sought (IBG, 2000). In China, biocontrol with a native fungal strain of C. gloeosporioides f. sp. cuscutae led to the commercialization of LuBao, in

1963, for the control of dodder (Cuscuta sp.) a parasitic weed in soybean. An improved formulation Lubao 2 is still in use today (Wan and Wang, 2001). Other potential uses of

Colletotrichum spp. as a mycoherbicide have been reported from many other regions of the world (Sharon, 1998; Nof et al., 1998; Gressel et al., 1998; Killgore et al., 1999).

Although many strains of Colletotrichum demonstrated adequate potential in field tests, they have not been developed as commercial products (Templeton, 1992b).

1.4.5. Mating systems in Colletotrichum spp.

Reproduction in most Colletotrichum spp. is vegetative. The sexual perfect stage has not been found in nature or in culture for many species. This is surprising given the

variability of members of the genus (Bryson et al., 1992). The unusual complex mating systems of Glomerella spp., ranging from self-fertile (homothallism), self-sterile but cross-fertile (heterothallism) and asexual strains may be correlated with its morphological instability (Bryson et al., 1992). Hence the genus Glomerella seems to be unique among the ascomycetes because a wide set of different mating types occur within strains and multiple mating types among the heterothallic strains occur (Hanau et al., 1998).

In Glomerella cingulata and G. graminicola both homothallic and heterothallic

strains exist (Vaillancourt et al., 2000). Homothallic strains of G. cingulata give rise to progeny with homothallic, heterothallic reproduction and also sterile progeny (Wheeler,

1954). G. graminicola mating systems include true heterothallism and also unbalanced heterothallism and true homothallism (Vaillancourt and Hanau, 1999; Vaillancourt et al.,

2000). Most homothallic and heterothallic strains are fertile and homothallic strains

frequently give rise to heterothallic progeny (Vaillancourt et al., 2000).

Extensive research in the 1940s and 1950s on the reproduction of G. cingulata

emphasized an unbalanced heterothallic mating system that resulted from mutation in the morphogenetic pathway necessary to homothallism (Chilton et al. 1944a, b; 1949a, b;

Edgerton et al., 1945; Wheeler, 1950; Hanau et al., 1998). In G. cingulata, a homothallic culture originated from a single ascospore produced asci containing four self-fertile and four self-sterile ascospores (Perkins, 1987). The new self-fertile ascospores germinate to form self-fertile colonies that give rise to asci containing once more four self-fertile and four self-sterile ascospores. Colonies originating from the self-sterile ascospores apparently mate with the self-fertile strains to form hybrid perithecia (Perkins, 1987). The

molecular basis of this unidirectional shift from self-fertile to self-sterile is not well understood (Glass and Kuldau, 1992).

Isolates of C. graminicola can be heterothallic and homothallic, but the homothallism may be a consequence of mutation and selection during repeated subculture and C. graminicola may be heterothallic in nature (Vaillancourt and Hanau,

1991). Conidial anastomosis between C. gossypii var. cephalosporioides isolates as well as sexual reproduction has been reported (Roca et al., 2004a). An experiment conducted with nitrate non-utilizing heterokaryon mutants from field isolates of C. destructivum, two C. gloeosporioides strains, C. gloeosporioides f. sp. jussiaea, C. fragariae, C.

malvarum and C. trifolium, demonstrated the self-compatibility between all the

heterokaryons except between C. destructivum and C. fragariae (Brooker et al., 1991).

1.4.6. Recombinant Colletotrichum spp.

Conidial anastomoses between isolates of two different Colletotrichum spp. (C.

lindemunthinum and C. gossypii) has been reported (Roca et al., 2004b). The growth of this hybrid was assessed to be less than the parental strains, however, in field conditions natural selection would ensure propagation of any hybrid with a selective advantage

(Roca et al., 2004b). Recombination between different lineages of C. graminicola from

maize and C. graminicola from Pooideae has also been suggested based on molecular

data (Crouch, 2006). Heterokaryons from C. gloeosporioides although exhibiting a slow

growth rate, were stable and could be perpetuated on minimal media (Chacko et al.,

1990). ‘The potential for sexual reproduction and gene flow within and between genetic subgroups of C. acutatum sensu lato and the influence on population structure remains largely unexplored’ (Guerber et al., 2003).

The commercialized mycoherbicide Collego®, formulated with C.

gloeosporioides f. sp. aeschynomene (pathogenic to northern jointvetch) can mate, in

vitro, with other strains of C. gloeosporioides pathogenic to different hosts [winged water

primrose, Ludwigia (Jussiaea) decurrens and pecan, Carya illinoensis] being capable of

producing viable, recombinant progeny with variable pathogenicity to apple fruits (Cisar

et al., 1994). Parental host specific strains from northern jointvetch and pecan crossed on the surface of northern jointvetch plants (in laboratory conditions), producing viable

progeny pathogenic only to apples (Cisar et al., 1996). Pecan and apple are both affected

by G. cingulata which causes pecan anthracnose and bitter-rot of apple (Rand, 1914).

Studies of heterokaryosis may indicate the potential for genetic exchange between related subspecies of Colletotrichum following field release of mycoherbicides (Chacko

et al. 1989, 1990). Vegetative compatibility among heterokaryon auxotrophic and nitrate

nonutilizing mutant isolates has been reported from C. orbiculare (Wasilwa et al., 1991),

Gromerella graminicola (Vaillancourt and Hanau, 1994) and from C. gloeosporioides f.

sp. aeschynomene (Chacko et al., 1990). In imperfect asexual stages of fungi, such as C.

gloeosporioides, heterokaryosis is likely to be a relevant factor in the rapid adaptation of

the pathogen to a new host (Sreenivasaprasad et al., 1993).

The environmental implications associated with gene flow between a genetically

engineered mycoherbicide and endemic fungi have not been determined (TeBeest and

Cisar, 1994; Cisar and TeBeest, 1995). Assessment of release risk for these agents must

include determination of the likelihood and consequences of gene flow between

bioherbicides and related fungi (Teng and Yang, 1993). Positive and negative aspects

must be balanced. Genetic manipulation may be useful to enhance the virulence of a

pathogen without a narrow host range or to reduce the host range of a highly virulent pathogen (Kistler, 1991; Wilson and Lindow, 1993).

1.4.7. Infection process

The pathogenicity of Colletotrichum spp. to plants is intrinsically linked with a sequence of a physiological and morphological chain of events (Dickman, 1998). After conidial attachment, germination and development of a germ tube, Colletotrichum spp.

penetrates the host through natural openings (stomata), wounds or by direct penetration

of the cuticle (Bailey et al., 1992). A highly specialized infection peg called an

appressorium allows them to penetrate a healthy host (Emmett and Parbery, 1975; Sutton,

1968; Dean, 1997). The appressorium has a turgor-based mechanical force that mediates

the direct penetration of a narrow peg through the host cuticule and epidermal layer aided

by the induction of extracellular hydrolitic enzymes that break the polymeric defenses of

the host (Money, 1995; Mendgen et al., 1996; Kolattukudy et al., 1998). After hyphal

penetration of an epidermal cell, intracellular infection vesicles that produce large

primary hyphae develop using nutrient assimilation within the plant tissue. These ramify

throughout the apoplast of the leaf tissue, producing secondary hyphae that move

throughout the plant via acervuli and are responsible for the shift from biotrophic to

necotrophic infection growth (Dickman, 1998; Wei et al., 2004). The pressure exerted at

the tip of the penetration peg of C. graminicola has been estimated at 17 micro Newtons

(µN) (Bechinger et al., 1999). To better visualize the exceptional force exerted by these

structures a 17µN pressure exerted by a human hand would allow it to support an 8-ton

school bus (Money, 1999a) or a whale (Money, 1999b). The pressure exerted by an

44 infection peg of the fungus Magnaporthe grisea has been estimated to be higher than any other living organism (Howard et al., 1991).

Colletotrichum spp. have different infection strategies, as follows: (1) intracellular hemibiotrophs, the fungus grows initially biotrophically and no symptoms are produced on the plant, followed by a visible destructive phase in which the pathogen becomes necrotrophic; (2) subcuticular intramural pathogens that are characterized by subcuticular growth that dissolves the pectic matrix of the epidermis cell walls, and (3) species exhibiting both hemibiotrophic and subcuticular intramural infections (Smith et al., 1999). Strict hemibiotrophs have non-overlapping biotrophic and necrotrophic fungal structures formed in different tissues of the plant. These hemibiotrophs are often found in

Colletotrichum spp., but rarely in other fungi such as Phytophthora infestants (Oliver and

Ipcho, 2004). The infection process of C. graminicola on different Poaceae hosts has been extensively evaluated. The following sequence of events was observed: spore germination occurs 2 h after inoculation, appressoria germinate within 6 h, cuticule penetration takes place 8 h after inoculation and infection hyphae develop inside the epidermal cells within 24 h (Perfect and Green, 2001; Khan and Hsiang, 2003; Wei et al.,

2004).

Fungal pathogenicity has been widely studied through different techniques (Gold et al., 2001). Different compounds and enzymes, produced by the host and pathogen,

Colletotrichum spp., have been isolated (Bailey et al., 1992; Dean, 1997; Kubo, 1998;

Prusky et al., 1998; Hutchison et al., 2000). Phytoplane microflora (Slade et al., 1986;

Jeffries, 1998), pH (Yakoby et al., 2000; Prusky et al., 2002), nitrogen sources (Drori et al., 2003) and genes expressed during infection have been studied and known to be

45 implicated in triggering appressoria formation, host infection and pathogenesis development (Takano et al., 2000; Dickman, 2004).

1.4.8. Quiescent infection by Colletotrichum spp.

The role of endophytic growth of fungi in plants has been associated with their symbiotic activity with the plant host. Endophytes may have antagonistic capabilities to other plant pathogens (i.e., fungi and bacteria) and insect plant pests, as well as, the ability to enhance the physiological activities of the plant host (e.g., drought tolerance and/or growth increase) (Azevedo et al., 2000).

Endophytic Colletotrichum spp. often penetrates leaves directly or enters through stomata openings and then ramifies to the mesophyll intercellularly, showing no symptoms of infection (Verhoeff, 1974; Carrol, 1995; Wei et al., 1997; Latunde-Dada,

2001). A physiological change, such as senescence of the host will trigger the necrotic phase of the fungus and the occurrence of visible necrosis. The quiescent (or latent) infection occurs in tropical and subtropical fruits in their early stages of development with symptoms appearing only in the near-mature or mature stage of fruit development

(Kuo, 1999). This occurs due to adverse physiological conditions temporarily imposed by the host (Prusly and Pumbley, 1992). Thus, appressoria do not germinate but the tissues are instead indirectly penetrated by vegetative growth through stomata (Latunde-Dada,

2001). After the fungus senses the substrate, a process associated with conidial adhesion and appressorium differentiation (Hoch, 2004) involving chemical signals from the host

(in its senescence or reproductive stage) will trigger the infection process through the germination of appressoria (Kolattukudy et al., 1998; Latunde-dada, 2001). It has been suggested that the lifestyle expressed by Colletotrichum spp. (mutualism, parasitism or

commensalism) is controlled by the host plant genotype (Redman et al., 2001). Also, it

has been reported that both chemical and physical stimuli between the fungus and plant

eventually alter gene expression, producing a cellular response (Memmott et al., 2002).

Experimental evidence reported by Kolattukudy et al. (1998) suggests that ‘conidia of C.

gloeosporioides are primed by a 2h contact with a hard surface that induces a set of

genes; this priming enables the conidia to respond to host signals such as host surface

wax and the host ripening hormone, ethylene, which induce another set of genes that

enables them to germinate and differentiate into appressoria’. These signals are released

from the host and pathogen and determine the type of symbiotic relationship that will

develop (Dickman 2000, 2004).

Methods to trace latent infection and gene expression by Colletotrichum spp. have

been developed. These are: DNA extraction and Polymerase chain reaction (Parikka and

Lemmetty, 2004) and green fluorescent protein-transgenic strains (Dumas et al., 2001;

Horowitz et al., 2002). Several Colletotrichum spp. have been reported to have

mutualistic or comensalistic lifestyles in plants. Colletotrichum spp. that have an

endophytic behavior are C. cocoodes, on tomato (Dillard and Cobb, 1998) and potato

(Tsor and Johnson, 2000), C. musae on banana and C. gloeosporioides on several plant

hosts such as cowpea and citrus (Zulfiqar et al., 1996; Cerkaukas, 1988; Latunce-Dada,

2001), C. demantium f. sp. truncatum, on soybean (Sinclair, 1991), C. gloeosporioides f.

sp. malvae on 10 different field crops (Makowski and Mortensen, 1998) and C. acutatum

on strawberry, pepper, egg-plant, etc. (Freeman et al., 2001b; Horowitz et al., 2002;

Zulfiqar et al., 1996).

Studies with Colletotrichum magna mutants showed that nonpathogenic life–

styles provide a broader host range than the parental pathogenic C. magna strains and that

nonpathogenic mutants conferred an array of mutualistic benefits on the host plant (e.g.,

resistance against wild type Colletotrichum spp. and drought, growth enhancement and biomass increase) (Redman et al., 2001).

1.4.9. Survival and dispersal under different abiotic conditions

Asymptomatic plants that are not considered hosts of C. acutatum, living

symbiotically or non-pathogenically, may serve as a potential focal source for infection.

These allow the pathogen to survive between growing seasons of the primary plant host

species (Freeman et al., 2001b).

Colletotrichum spp. generally produces sticky masses of spores in acervuli

(TeBeest, 1991). Insect vectors transmit viable conidia of Colletotrichum spp. to suitable

hosts. Chrysomellid beetles (Chrysolina hyperici Forest.) can enhance the biocontrol of

St. Johnswort (Hypericum perforatum L.) seedlings by transmitting C. gloeosporioides f.

sp. hypericum during foraging and feeding (Morrison et al., 1998). Grasshoppers

(Conocephalus fasciatus and Melanophus differentialis) and green treefrogs (Hyla

cinerea) may be important vectors of the fungus in northern jointvetch in rice (Yang and

TeBeest, 1992, 1994).

It has been determined that conidia of C. acutatum in an infected leaf showed a

decreased survival rate under moist or saturated conditions and increased survival

(maintaining pathogenic potential) under dry conditions, living up to 12 months after

being incorporated into sandy soil with a pH of 6.9 (Norman and Strandberg, 1997).

Overwintering survival of C. acutatum from non-mummified strawberry fruits located on

and below the soil surface have been also reported (Wilson et al., 1992) as well as in high bush blueberry, Vaccinium corymbosm (DeMarsay and Oudemans, 2001). C. acutatum

conidial spores are more tolerant to extended periods of low temperatures than C.

gloeosporioides, where spore germination, survival and appressoria formation can be compromised (Everett, 2003). Survival in the soil may be enhanced by low temperatures or greater burial depths (Eastburn and Gubler, 1990). Summer survival of C. acutatum

has been determined to be lower (4%) than C. gloeosporioides (96%) after soil burial of

infected strawberry crowns (Ureña-Padillo et al., 2001). Colletotrichum acutatum was

not recovered after 56 d of summer burial; C. gloeosporioides recovery was 10-20% after

70 d, suggesting that oversummering crop debris did not serve as inoculum for epidemics of Colletotrichum spp. crown rot (Ureña-Padillo et al., 2001).

Colletotrichum coccodes survives longer as free sclerotia in the soil than

associated with plant tissue. After eight yr the viability of free sclerotia was 0.0, 90 and

80% at the soil surface, 10 and 20 cm soil depth, respectively (Dillard and Cobb, 1998).

This may result from greater fluctuation in temperature and moisture at the soil surface

(Dillard and Cobb, 1998). Sporogenic germination of sclerotia originated from C. cocoodes, occurred at a wide range of temperatures, with production of conidial masses from 10-34o C in aerated plates, in vitro (Sanogo and Pennypacker, 1997).

Sclerotium are important dormant or quiescent structures of many plant

pathogens, being extremely resistant to harsh conditions (i.e., desiccation, low

temperatures, fungicides, etc.), enabling fungal pathogens to overcome adverse periods

and germinate when conditions improve (Coley-Smith and Cooke, 1971). They originate

initially from a cluster of hyphae - a mass of tightly interwoven hyphal cells - then

49 differentiate into external thick-walled and melanized hyphae and internal extended vacuolated hyphae. The inner tissue is nutrient rich with abundant lipid and glycogen reserves surrounded by a cortex of cells with thicker walls (Tu, 1980; Carlisle and

Watkinson, 1997).

1.5. Research objectives

The present research intends to contribute to a better understanding of the etiology of fungal epizootics in general and of Colletotrichum in particular, through the study of the natural occurring epizootic in populations of F. externa in the northeastern hemlock forests. The main objectives of this research are:

1. To identify the causal agents of the emergent epizootic within populations of the

northeastern elongate hemlock scale, Fiorinia externa, by molecular and

morphological approaches, as well as its geographic distribution;

2. To characterize morphologically, biologically and phylogenetically the

Colletotrichum sp. strain commonly isolated from areas of the epizootic;

3. To assess the virulence of the ubiquitous Colletotrichum sp. strain present in the

geographic range of the epizootic to four families of insects and five families of

plants.

Table 1.1. Reports of fungal species recovered from populations of scale insects

Fungi Host scale Location Source

Aschersonia turbinata Berk. Wax scale (Ceroplastes floridensis Comstock) Florida (USA) a , b

Aschersonia cubensis Berk. and Pyriform scale (Pulvinaria pyriformis Ckll.) Florida (USA) b

Curt. Liriodendron scale [(Toumeyella liriodendri (Gmelin.)] Puerto Rico (USA)

Tessellated scale (Eucalymnatus tessellates)

Aschersonia aleyrodis (Webber) West Indian Red scale [Selenaspidus articulatus (Morgan)] Brazil p

Parlatoria ziziphus (Lucas)

Aschersonia basicystis Berk. & Unidentified Coccidae Venezuela q

MA Curtis.

Aschersonia placenta Berk. and Asterolecanium ungulata Russel Malaysia r , s

Br. India t

Aschersonia duplex Berk. Metaceronema japonica Mask. China v

Hypocrella duplex (Berk.) Coccoidea Caucasus y

Petch

Aschersonia sp. Japanese scale (Lopholeucapsis japonica Cockerell) Georgia (Caucasus) u

Brown scale [Saissetia coffeae (Walker)] India w

Hypocrella turbinata (Berk.) Wax scale (Ceroplastes floridensis Comstock) Florida (USA) d

Petch Purple scale [Lepidosaphes beckii (Newm.)] Puerto Rico

Soft brown scale (Coccus hesperidum Linn.) Costa Rica

Hypocrella javanica (Penz. and California red scale (Aonidiella aurantii (Mask.) Japan

Sacc.) Unidentified scales Ceylon

India

Hypocrella reinekiana P. Henn. Lecanium sp. Formosa

Parlatoria sp. Ceylon

Hypocrella raciborskii Asterolecanium ungulata Russel Malaysia r , s

Hypocrella palmae(Berk. & Coccoidea Black sea coast y

MA Curtis) Sacc.

Calonectria decora (Wallr.) Coccoidea Caucasus

Sacc

(Continue)

Table 1.1 (Continued). Reports of fungal species recovered from populations of scale insects

Fungi Host scale Location Source

Hirsutella besseyi Fisher Coccoidea (Citrus trees) Florida (USA) x

Hirsutella lecaniicola (Jaap) Parthenolecanium corni Bouché Eastern Russia z

Petch

Hymenostilbe lecaniicola Coccoidea USA x

(Jaap) Mains Canada

Romenia

Germany

Soviet Union

Myriangium duriaei Mont. Ischnaspis filiformis Douglas Java e

Aspidiotus camelliae Signoret Ceylon e

Chionaspis biclavis Maskell

San José Scale (Quadraspidiotus perniciosus Comstock) Florida (USA) f

Long scale (Lepidosaphes gloverii Pack) Florida (USA) b

Purple scale (Lepidosaphes beckii (Newm.)

Chaff scale (Parlatoria pergandii Comst.)

Wide range of species USA d

Central America

South America

Ceylon

Philippines

Myriangium floridanum Snow scale (Chionaspis citri Comst.) Florida (USA) d

Hoehn. Purple scale (Lepidosaphes beckii (Newm.) India x

Africa

Myriangium montagnei Berk. Purple scale (Lepidosaphes beckii (Newm.) New Zealand d

Tasmania

Australia

Myriangium asterinosporum Coccoidea Canada x

(Ellis & Everh.) Eastern USA

(Continue)

Table 1.1 (Continued). Reports of fungal species recovered from populations of scale insects

Fungi Host scale Location Source

S. coccophila Tul. Lecanium sp. British West Indies g

Ophionectria coccicola E.& E. Purple scale (Mytilaspis citricola Pack.) Islands

Myriangium duriaei Mont Aspidiotus articulatus Morgan

Lecanium mangiferae Green

Chephalosporium lecanii Diaspis sp. British West Indies g

Zimmermann Islands

Sphaerostilbe coccophila Tul. San José Scale (Quadraspidiotus perniciosus Comstock) Florida (USA) h

Japan i

Purple scale [Lepidosaphes beckii (Newm.)] Florida (USA) b

Chaff scale (Parlatoria pergandii Comst.)

Sphaerostilbe flammea Tul. Snow scale (Chionaspis citri Comst.) Florida c

Panama

Australia

Cephalosporium lecanii Zimm. Pyriform scale [Protopulvinaria pyriformis (Ckll.)] Florida (USA) f

Hemispherical scale [Saissetia hemisphaerica (Targ.)]

Wide range of hosts West Indies Islands

Ceylon

Puerto Rico

Cephalosporium lecanii Zimm. Wide range of hosts British Guiana f

Brazil

New Zealand

Cephalosporium acremonium Coccoidea Germany x

Corda Austria

Netherlands

England

Italy

Romenia

Soviet Union

(Continue)

Table 1.1 (Continued). Reports of fungal species recovered from populations of scale insects

Fungi Host scale Location Source

Cephalosporium sp. Pulvinaria sp. Argentina f

Brown scale [Parthenolecanium corni (Bouché)] Italy

Podonectria sp. Diaspidid scales South Africa l

Sphaerostilbe aurantiicola

(Kerk. and Br.)

Podonectria aurantii (V. Höh) Citrus scales Brazil m

Petch Black parlatoria scale [(Parlatoria zizyphus (Lucas)] Formosa

Podonectria coccicola (Ellis Chaff scale (Parlatoria pergandii Comst.) Florida (USA) d

and Everh.) Petch Purple scale [Lepidosaphes beckii (Newm.)] Japan

Glover’s scale [Lepidosaphes gloverii (Pack.)] South America

Barbados

Java

Citrus scales India

Ophionectria coccicola E. and Long scale (Lepidosaphes gloverii Pack.) Florida (USA) m

E. Purple scale [Lepidosaphes beckii (Newm.)]

Chaff scale (Parlatoria pergandii Comst.)

Spicaria sp. Cottony cushion scale (Icerya purchasi Maskell) Puerto Rico n

Nectria diploa (B. and C.) Purple scale [Lepidosaphes beckii (Newm.)] Florida (USA) f

Long scale (Lepidosaphes gloverii Pack.) Australia

Black scale [Saissetia oleae (Bern.)] Formosa

California Red scale [Aonidiella aurantii (Mask.)] Central America

South America

West Africa

Neozygites lecanii Lecanium viridae Green Poland k

(Zimmermann)

Beauveria sp. Purple scale [Lepidosaphes beckii (Newm.)] Asia d

Aspergillus sp. Brown scale [Parthenolecanium corni (Bouché)] Florida (USA)

Verticillium sp. Glover’s scale [Lepidosaphes gloverii (Pack.)] Italy

Fusarium sp. Black Thread scale [Ischnaspis longirostris (Sign.)] England

(Continue)

Table 1.1 (Continued). Reports of fungal species recovered from populations of scale insects

Fungi Host scale Location Source

Beauveria sp. California red scale (Aonidiella aurantii Comst.) Australia d

Aspergillus sp. Aspidiotus sp. Algeria

Verticillium sp. Oystershell scale Argentina

Fusarium sp. Dictyospermum scale South Africa

Ceylon

Fusarium coccophilum (Desm.) Coccoidea Caucasus y

Wollenweber & Reinking

Fusarium microcera Bilai Coccoidea Soviet Union y

[telemorph: Calonectria decora

(Wallr.) Sacc.]

Torrubiella lecanii Johnston Hemispherical scale [Saissetia hemisphaerica (Targ.)] Cuba d

Puerto Rico

Pegiotrichum saccarinum Raw. Unidentified scale Brazil

Acrostalagmus albus Pr. Soft brown scale (Coccus hesperidum Linn.) Italy

Isaria sp. Black scale [Saissetia oleae (Bern.)] North Carolina o

Virginia

Sources: a – Webber (1894, 1897) n – Wolcott (1940) b – Berger (1921) o – Charles (1941) c – Fawcett (1944) p - Gravena et al. (1988, 1992) d – Fawcett (1948) q - Rojas (2000) e – Parkin (1906) r - Lim et al. (1991) f – Watson and Berger (1937) s - Ibrahim and Kon (1992) g – South (1910) t - Devnath (1986) h –Rolfs (1897) u - Shiryaeva (2000) i – Rolfs and Fawcett (1908) v - Zhuan (1988) j – Kuwana et al. (1904) w - Das and Ganguli (1961) k – Balazy (1999) x - Evlachova (1974) l – Annecke (1963) y – Koval (1974) m – Rao and Sohi (1980) z – Borisov et al. (2001)

Table 1.2. Reports of fungi recovered from Fiorinia species

Fungi Host Scale Location Source

Sphaerostilbe coccophilus F. fioriniae Ceylon a

Nectria diploa Berk. and Curt. F. rubrolineata Ceylon b

Pyrenochaeta sparsibarba F. juniperii Ceylon b

Muricularia calva

Stilbum coccorum

Fusarium sp. F. thea India c

Sources: a – Parkin (1906) b – Petch (1922) c – Das and Das (1962)

Table 1.3. Sequence of morphological identifications of the most commonly isolated fungus from diseased F. externa in the states of Connecticut, New York, New Jersey and Pennsylvania.

Year Institution Identifier ID

2003 Entomology Research Laboratory, Drs. Svetlana Gouli and Hypocrella sp.

University of Vermont, USA Vladimir Gouli

2005 Mycological Society of China, China Dr. Zengzhi Li Aschersonia

marginata

2005 Cornell University, USA Dr. R. Humber Fusarium merismoides

Systematic Botany and Mycology Laboratory Dr. A. Rossman

2005 Cornell University, USA Dr. Kathie T. Hodge Tubercularia sp.

Cornell’s Plant Pathology Herbarium

2006 National Center for Genetic Engeneering Dr. Nigel Hywel-Jones Myriangium sp.

and Biotechnology, Thailand

CHAPTER 2

Fungi Associated with a Natural Epizootic in Fiorinia externa Ferris (Hemiptera: Diaspididae) populations

José A. P. Marcelino, Svetlana Gouli, Rosanna Giordano, Vladimir V. Gouli, Bruce L. Parker, Margaret Skinner

Abstract: Stands of eastern hemlock in the northeastern United States are in decline, in part from the activity of elongate hemlock scale, Fiorinia externa Ferris (Hemiptera: Diaspididae). From 2001 to the present a natural epizootic has been observed in populations of F. externa. Initially discovered at the Mianus River Gorge Preserve in Bedford, NY, the epizootic also has been detected in Pennsylvania, New Jersey and Connecticut. Understanding and assessing the identity of the pathogenic microorganisms responsible for this natural mortality is crucial for developing biological controls for this pest. We have isolated and taxonomically and genetically identified entomopathogens, phytopathogens and endophytic fungi associated with F. externa. Isolates of the following were obtained: Colletotrichum sp., Lecanicillium lecanii, Beauveria bassiana, Cordyceps sp. anamorph, Myriangium sp., Mycosphaerella sp. anamorph, Nectria sp., Botrytis sp., Phialophora sp. and Fusarium sp.

Keywords: epizootic, biological control, entomopathogenic fungi, Tsuga canadensis

______

José A. P. Marcelino, Entomology Research Laboratory, 661 Spear St., University of Vermont, Burlington, VT, 05405 – Phone: (802) 656 5441; Fax: (802) 656 5441; [email protected]. Svetlana Gouli, Entomology Research Laboratory, 661 Spear St., University of Vermont, Burlington, VT, 05405 – Phone: (802) 656 5438; Fax: (802) 656 5441; [email protected]. Rosanna Giordano, Illinois Natural History Survey, Division of Biodiversity and Ecological Entomology, 1816 S. Oak St., Champaign, Illinois, 61820; - Phone: (217) 333 8627; Fax: (217) 244 0729; [email protected]. Vladimir V. Gouli, Entomology Research Laboratory, 661 Spear St., University of Vermont, Burlington, VT, 05405 – Phone: (802) 656 5438; Fax: (802) 656 5441; [email protected]. Bruce L. Parker, Entomology Research Laboratory, 661 Spear St., University of Vermont, Burlington, VT, 05405 – Phone: (802) 656 5440; Fax: (802) 656 5441; [email protected]. Margaret Skinner, Entomology Research Laboratory, 661 Spear St., University of Vermont, Burlington, VT, 05405 – Phone: (802) 656 5440; Fax: (802) 656 5441; [email protected].

Acknowledgments: We thank Lora Schwarzberg (New York State Department of Environmental Conservation) and Teri Hata (University of Vermont, Entomology Research Laboratory) for valuable help collecting specimens and for technical assistance. We also thank Drs. David TeBeest (University of Arkansas) and Roberto Cesnik (EMBRAPA, Brazil) for their scientific input and insights. This work was funded in part through a grant awarded by the Northeastern Area State and Private Forestry, USDA Forest Service (# 04-CA-11244225286) and is in partial fulfillment of requirements for the PhD degree of J.M. at the University of Vermont. ______

VIDENCE OF SIGNIFICANT DECLINE AND DEATH of eastern hemlocks [Tsuga E canadensis (L.) Carrière] has been observed in the northeastern United States. It has been attributed in part to feeding by the elongate hemlock scale, Fiorinia externa

Ferris (Hemiptera: Diaspididae) and the hemlock woolly adelgid, Adelges tsugae Annand

(Hemiptera: Adelgidae) (Colbert et al. 2002, Danoff-Burg and Bird 2002). Fiorinia

externa is found along the eastern seaboard from Massachusetts to North Carolina.

Populations reach as far west as Ohio, with the highest density of infestation occurring

within a 300 km radius of New York City (USDA Forest Service 2004). Natural

biological control agents have not succeeded in keeping populations of F. externa below

damaging levels. Insecticide treatment of this pest has also proved ineffective as

populations resurge rapidly after application (McClure and Fergione 1977). High

parasitism rates of F. externa by the aphelinid parasitoid, Encarsia citrina Craw

(Aspidiotiphagus citrinus Howard) (Hymenoptera: Aphelinidae) have been observed in

its native region of Japan (McClure 1986), but this species does not occur in the

northeastern United States (McClure 1977). The armored shield of F. externa helps to protect it from insecticides, natural enemies and adverse weather conditions.

Given the rapid decline of hemlock, it is imperative that control methods are devised that are ecologically sound and cost effective. In 2001 an epizootic within populations of F. externa was discovered in the Mianus River Gorge Preserve in Bedford,

NY, where 13-27% of sampled insects were covered by profuse sclerotic masses which often concealed the insects’ body (McClure 2002). Forested areas with similarly infected scales have been found in several counties in New York, Pennsylvania, Connecticut and

New Jersey (Parker et al. 2005).

To evaluate the role of epizootics in the suppression of insect pests, it is essential

to determine the causal agent(s) involved in the disease. The objective of this study was

to morphologically characterize and molecularly identify the mycobiota associated with

the epizootic of F. externa.

Materials and Methods

Sampling

Thirty-six sites were surveyed and collections of infected F. externa adults (i.e., with sclerotic masses or other visible signs of mycoses) were made from 2004 to 2006 in

New York (nine counties), Pennsylvania (three counties), Connecticut (two counties) and

New Jersey (one county) (Figure 2.1). Surveys were made in areas with a high percentage

of hemlock trees, based on land cover maps from the NY Dept. of Environmental

Conservation, and in hemlock stands in state and county parks, camping areas, city-

owned reservoirs, forest preserves, sanctuaries, historic sites and arboretums. In each

hemlock stand, a minimum of 100 hemlocks were randomly sampled, taking five 40-cm

long branches per tree, from which to obtain infected scales and isolate fungi. In 10 sites

in New York and four in Pennsylvania, a sub-sample of 10 twigs from five branches was

used to make a visual estimate of the percentage of F. externa showing evidence of

fungal infection, i.e., sclerotic masses on the scale surface, abnormal pigmentation and

loss of turgor or mycelium arising from the body of the insect.

NH NY

Legend: Initial epizootic report (2002) Sample sites with diseased Fiorinia externa (2002-2006)

Km 0 30 60 120 180 240 1 centimeter equals 34.964 kilometers

Figure 2.1. Geographical distribution of sites with diseased Fiorinia externa populations (2002-2006).

Isolation of fungi

Scale-infested twigs were inspected under a stereomicroscope, and those with

signs of mycosis were surface sterilized by dipping in a solution of 2.5% sodium

hypochlorite with 0.1% Silwet L-77 for 45 s, rinsed with sterile distilled water (SDW)

and placed on potato dextrose agar (PDA) supplemented with Penicillin (5 ml/l) and

Streptomycin (12.5 ml/l). Other selective media were also used for isolation of fungi:

Sabouraud dextrose agar and yeast, Sabouraud dextrose agar supplemented with egg yolk

and milk, bacto-agar with beef extract, bacto-agar with liver extract, and modified media

supplemented with Schneider’s insect medium (Sigma Chemical Co., St. Louis, MO). All

isolates were purified using standard dilution techniques (Schmitthenner and Hilty 1962),

from which monosporic lines were generated for the most commonly retrieved fungi,

using a stage-mounted DC3001 micromanipulator (World Precision Instruments, Inc.,

Sarasota, FL).

Molecular identification of fungi

DNA of the most commonly isolated fungi was extracted from one-week-old

cultures of the fungi isolated from the pest, from sclerotia and also F. externa enclosed in sclerotia (Table 2.1).

Table 2.1. Pure fungal isolates recovered from Fiorinia externa

DNA GenBank accession numbers Year Reference Fungal group Species/Genus Geographic origin Extraction Collected code 28S ITS from

Entomopathogens Myriangium sp. Bayberry Lane, NY * 2006 CEHS208 1. Culture EF464574 EF464585

(N= 62 isolates) (Σ= 121) Litchfield, CT SEHS208 2. Sclerotia EF464575 EF464586

Litchfield, CT SINEHS208 3. Sclerotia EF464576 EF464587

+ F. externa

Cordyceps sp. anamorph 2005 CEHS133a 1. Culture EF464571 EF464589 Valley Forge, PA (N= 41 isolates) INEHS133a 2. F. externa EF464572 EF464583 63

L. lecanii Mount Tom Forest Preserve, 2005 EHS132 1. Culture EF464573 EF464584

(N= 2 isolates) MA

B. bassiana Mount Tom Forest Preserve, 2005 EHS163 1. Culture □ □

(N= 16 isolates) MA

Phytopathogens Mycosphaerella sp. Pawling Preserve, NY 2006 EHS201 1. Culture EF619924 EF619925

(Σ= 58) anamorph

(N=1 isolate)

♦ Colletotrichum sp. NY; PA; NJ; CT 2004/05 EHS48 1. Culture EF464580 EF464593

(N= 54 isolates)

(Continue) 63

Table 2.1 (Continued). Pure fungal isolates recovered from Fiorinia externa

DNA Year Reference GenBank accession numbers Fungal group Species/Genus Geographic origin Extraction Collected code from 28S ITS

Phytopathogens Nectria sp. Valley Forge, PA 2006 EHS144 1. Culture EF464570 EF464588

(Σ= 58) (N= 1 isolate)

Botrytis sp. Bayberry Lane, NY 2006 EHS265 1. Culture □ □

(N= 1 isolate)

Fusarium sp. South Salem, NY 2006 EHS290 1. Culture □ □ 64 (N= 1 isolates)

Endophyte Phialophora sp. South Salem, NY 2006 EHS 291 1. Culture EF464577 EF464590

(N= 1 isolate) (Σ= 1)

♦ Counties: New York (Columbia, Dutchess, Kings, Orange, Putnam, Queens, Rockland, Ulster and Westchester); Pennsylvania (Bucks, Chester

and Northampton); New Jersey (Passaic); Connecticut (Fairfield and Litchfield).

* Origin of samples for use in DNA extraction.

□ Only morphological identification was performed.

The Power SoilTM DNA kit (Mo Bio Laboratories, Inc., Carlsbad, CA) was used

according to the manufacturers’ instructions with the following exceptions: 1) Samples

were shaken for 5 min at 5.5 m/s to facilitate opening of the fungal cell walls using a

FastPrepTM FP120 machine (Thermo Savant, Holbrook, NY); and 2) DNA was eluted using 100 µl of diluted elution buffer (1:15) (Quiagen, Valencia, CA) and concentrated to

20 µl with a speed vacuum (Eppendorf Centrifuge 5415C, Vaudaux, Switzerland) prior to

downstream applications. Polymerase Chain Reaction (PCR) was performed using

Ready-To-Go RT-PCR beads (Amersham Biosciences Inc., Piscataway, NJ). The D1/D2

region of the 28S ribosomal DNA commonly used for phylogenetic analysis at the genus

level and above (Hillis and Dixon 1991) was amplified with primers NL1 and NL4

(O’Donnell 1992, 1993).

The internal transcribed spacers (ITS) variable region was amplified using primers

ITS1 and ITS4 (White et al. 1990) for within-species differentiation (Talhinhas et al.

2002, Afanador-Kafuri et al. 2003). The 28S ribosomal DNA gene was amplified using

the following protocol: initial denaturation at 95o C for 2 min followed by 30 cycles of

95o C for 30 s, annealing at 50o C for 30 s and elongation at 72o C for 1 min. The protocol

used for the amplification of the ITS region was the same as above, except that the

annealing temperature was raised to 52o C. PCR products were purified using the

QIAquick PCR purification kit (Qiagen, Valencia, CA) or Centri SpinTM columns

(Princeton Separation, Adelphia, NJ). DNA was stored at 4o C. PCR products were

sequenced using BigDye v1 (Applied Biosystems, Foster City, CA) according to the

following protocol: initial denaturation at 95o C for 3 min followed by 30 cycles of 95o C

for 10 s, annealing at 50o C for 5 s, and elongation at 60o C for 2 min.

Sequencing reactions were run on a 3130xl Genetic Analyzer (Applied

Biosystem, Foster City, CA). Chromatograms were examined, and contiguous sequences

were generated, using SequencherTM (Gene Codes Corporation, Ann Arbor, MI).

Sequences generated from this study were compared to related sequences in GenBank

using BLAST (Altschul et al. 1990). Matches obtained from these searches were checked

against our preliminary morphological identifications. Sequences obtained for this study

were deposited in GenBank (accession numbers included in Table 2.1).

Morphological identification

Pure cultures of fungi from F. externa were prepared for microscopic

morphological identification using the methods of Humber (1997) and contrasted with reference taxonomical guides (Carmichael et al. 1980, Barnett and Hunter 1998, Samson et al. 1988). Scales with superficial signs of mycosis were also prepared for

morphological identification of fungal propagules using scotch tape impressions

according to the method of Gouli et al. (2005). All fungal isolates were deposited in the

entomopathogenic fungal germplasm collection at the Entomological Research

Laboratory, University of Vermont, Burlington, VT, USA.

Results and Discussion

In the site where the epizootic was first reported up to 36.8% of the sampled

scales were partially or completely covered with sclerotia in 2004. These sclerotia were

usually brown, irregular masses when associated with the new generation of infected

scales and turned black after several months (Figure 2.2).

Figure 2.2. Fiorinia externa showing typical symptoms of fungal infection, and representative propagules from several entomopathogenic fungi found associated with F. externa in epizootic areas. A. Scale covered with a fresh fungal sclerotium; B. Old, weathered sclerotia covering scale adults; C. Myriangium sp. hyphae (CEHS208); D. Beauveria bassiana mycosis; E. Beauveria bassiana conidia (EHS163) F.

Lecanicillium lecanii (EHS132) mycosis; G. Lecanicillium lecanii conidia; H. Cordyceps sp. anamorph

mycosis; I. Cordyceps anamorph sp. conidia (CEHS133a); J. Colletotrichum sp. conidia (EHS48); K.

Fusarium sp. conidia (EHS290); L. Mycosphaerella anamorph sp. conidia (EHS201). Bars: A, B, D, F, H= 0.5 mm; E, G, L = 40 µm; J, K, I, C = 25 µm.

No evidence of fungal hyphae penetrating hemlock needles was detected. The

proportion of scales showing signs of fungal infection varied among the different sites.

An approximate estimate of the percentage of scales with sclerotic masses in the sites sampled within New York, based on the number of infected scales in the sample stems

(10 hemlock twigs/5 branches) averaged ca. 26%, ranging from 61.8% in South Salem, to

2% in Esopus. In Pennsylvania, the proportion of infected scales on the sampled twigs was 9.5% ranging from 27.6 % at the Ralph Stover State Park to 0.8% in Jacobsburg. A variety of fungi were cultured and microscopic observations of conidia, conidiophores, mycelium and appressoria made. Molecular analysis using the 28S and ITS sequences facilitated the identification of the genera of the fungi. In many cases molecular analyses confirmed morphological identifications. A total of 180 pure fungal cultures were obtained from infected F. externa adults, comprising four entomopathogens, five phytopathogens and one endophytic species. Myriangium sp. was molecularly identified with DNA extracted from a pure fungal culture and also from single sclerotia, with no insect body parts, as well as sclerotia with enclosed mummified F. externa. Cordyceps sp. was molecularly identified with DNA extracted from both a pure fungal culture and F. externa adults with signs of mycosis. For all other fungi identified molecularly, DNA was obtained from pure fungal cultures. A total of 20 molecular identifications were obtained from F. externa with signs or symptoms of disease (Table 2.1).

Entomopathogenic Fungi

A total of 121 fungal isolates known as entomopathogenic species were collected comprising 16 isolates of Beauveria bassiana, two Lecanicillium lecanii, 41 Cordyceps sp. anamorph and 62 isolates of Myriangium sp.

The entomopathogenic isolates were among the most well-known and

commercially available fungi. For example, B. bassiana is known to be pathogenic to 100 insect species (McCoy et al. 1988); Lecanicillium lecanii has been used to control coccids, aphids and whiteflies (Feng et al. 2000). Cordyceps is primarily a parasite of insects (Nikoh and Fukatsu 2000), while Myriangium has proven effective in eradicating populations of scales (Van Epenhuijsen et al. 2000).

Myriangium sp. was the most prevalent entomopathogenic fungus isolated from

F. externa. It was isolated from fresh and old weathered single sclerotia (i.e., portion of sclerotia with no insect body parts) and from sclerotia enclosing F. externa. This fungus

was not recovered from all of the sample sites, which may be due to the difficulty in isolating this organism, rather than non-occurrence. Myriangium spp. are usually

exclusive pathogens of scale insects and form a stromata identical to the one present in all

the epizootic sites sampled (Miller 1940). Earlier studies described the presence of

Myriangium spp. on scale insects in Florida but its existence in latitudes other than

tropical and sub-tropical is unusual (Fisher et al. 1949). The use of Myriangium spp. for

management of scale insects has been limited due to the difficulties associated with its

culture in vitro. Its host specificity makes it an optimal candidate for management of F. externa.

Phytopathogenic and Endophytic Fungi

Fifty nine phytopathogenic isolates were recovered from F. externa: 54 isolates of

Colletotrichum sp., two Botrytis sp., one Nectria sp., one Mycosphaerella sp. anamorph and one Fusarium sp. One endophytic fungal isolate of Phialophora sp. was also recovered (Table 2.1).

Given the mode of action of phytopathogenic fungi, it is possible that these

isolations are a result of secondary infections in host insects by other pathogens that may

have overtaken the immune system of the host. Colletotrichum sp., a well-known phytopathogen, was the most prevalent fungus associated with the F. externa epizootic and was present in all localities sampled. In addition, this strain was topically applied to

F. externa in controlled bioassays and recovered after surface sterilization, confirming

Koch’s postulates. The above suggests that this Colletotrichum strain could have a key role in the epizootic. This genus has been widely associated with phytopathogenicity

(Latunde-Dada 2001), not entomopathogenicity. However, C. gloeosporioides, has been

reported to infect the scale Orthezia praelonga Douglas, (Hemiptera: Ortheziidae) on

citrus in Brazil (Cesnik and Ferraz 2000) where it is currently under development for

commercialization as a myco-pesticide (R. Cesnik pers. comm.). Due to the

phytopathogenic activity of this genus, it is critical to evaluate the agent-host-plant

interactions (Sands and Van Driesche 2000), and to conduct a comprehensive assessment

of its biology and genetics. A detailed molecular characterization of this strain is

underway to further evaluate its relationship with other phytopathogenic isolates.

Colletotrichum sp. from the epizootic areas has been found to have an endophytic

association with many locally occurring plants (Marcelino et al. submitted). We

hypothesize that due to its ubiquitous presence, the Colletotrichum sp. isolated plays a

significant role in the dynamics of the epizootic in populations of F. externa.

The Fungal Complex

It is possible that more than one fungal species is responsible for this epizootic.

The virulence and infectivity of pathogens are known primary factors involved in the

70 initiation and development of epizootics. It is common for epizootiological studies to only consider one pathogen infecting an insect population, though an insect can be affected simultaneously by multiple pathogenic species (Fuxa and Tanada 1987, Tanada and Kaya 1993). Several fungi have been implicated in the wide dissemination of mycoses of several species of scale insects in citrus orchards (Watson and Berger 1937,

Fawcett 1948, McCoy 1985), especially the fungal genera Aschersonia, Aegerita,

Verticilium, Sphaerostilbe, Podonectria, Myriangium and Hirsutella (Lord 2005). A complex of entomophthoralean fungi has been reported to infect other hemipteran insects

(Barta and Cagan 2003). However, there are no data showing synergistic effects of pathogenic agents on mortality or spread of disease. Future research should focus on the virulence and infectivity of individual pathogens associated with F. externa. Equally as important is to determine the combined effects of multiple infections as these may accelerate the progress of disease and the spread of this epizootic. The fungi described in this paper associated with F. externa may be functioning as a natural regulatory factor in populations of this pest. Enhancing the impact of this complex of fungi through augmentative techniques could help to suppress F. externa populations in areas where the epizootic has not reached. This control could be achieved with low production costs and minimal environmental safety concerns.

Understanding the patterns and processes of the epizootic in F. externa populations in the Northeast will provide opportunities for the management of this pest in the future.

Moreover, a better understanding of the possible synergistic actions among the fungi identified may elucidate the etiology of epizootics and fungal diseases in other insects.

The entomopathogenic fungal species isolated from F. externa adults may prove to be

71 good candidates for the management of other hemipteran pests of hemlock forests, i.e., the hemlock woolly adelgid.

Literature Cited

AFANADOR-KAFURI, L., D. MINZ, M. MAYMON, AND S. FREEMAN. 2003. Characterization of Colletotrichum isolates from tamarillo, passiflora, and mango in Colombia and identification of a unique species from the genus. Phytopathology 93:579–587. ALTSCHUL, S.F., W. GISH, W. MILLER, E.W. MYERS, AND D.J. LIPMAN. 1990. Basic local alignment search tool. J. Mol. Biol. 215:403–410. BARNETT, H. L., and B.B. HUNTER. 1998. Illustrated genera of imperfect fungi. Fourth edition. APS Press, St. Paul, MN. 218 p. BARTA, M., AND L. CAGAN. 2003. Entomophthoralean fungi associated with the common nettle aphid (Microlophium carnosum Buckton) and the potential role of nettle patches as reservoirs for the pathogens in landscape. J. Pest Sci. 76 (1):6–13. CARMICHAEL, J.W., KENDRICK, W. B., I.L. CONNERS, AND L. SIGLER. 1980. Genera of hyphomycetes. The University of Alberta Press. 369 p. CESNIK, R., AND J.N.G. FERRAZ. 2000. Orthezia praelonga, 1891 (Hemiptera, Ortheziidae) biologia, controle quimico e biologico. Embrapa Meio Ambiente, Jaguariúna, Brazil, Boletim de Pesquisa 9. 27 p. (in Portuguese) COLBERT, J., M. SEESE, AND B. ONKEN. 2002. Hemlock Woolly Adelgid impact assessment survey. P. 323-328 in Proc. of the hemlock woolly adelgid in the eastern United States symposium, Onken, B., R. Reardon, and J. Lashomb (eds.). New Jersey Agricultural Experiment Station, Rutgers University, New Brunswick, NJ. DANOFF-BURG, J.A., AND S. BIRD. 2002. Hemlock woolly adelgid and elongate hemlock scale: Partners in crime? P: 254-268 in Proc. of the hemlock woolly adelgid in the eastern United States symposium, Onken, B., R. Reardon, and J. Lashomb (eds.). New Jersey Agricultural Experiment Station, Rutgers University, New Brunswick, NJ. FAWCETT, H.S. 1948. Biological control of citrus insects by parasitic fungi and bacteria. P. 627-664 in The citrus industry, Vol. 2, Batchelor, L. D., and H. J. Webber (eds.), University of California Press, Berkeley, CA. FENG, K.C., B.L. LIU, AND Y.M. TZENG. 2000. Verticillium lecanii spore production in solid-state and liquid-state fermentations. Bioprocess Biosyst. Eng. 23:25-29. FISHER, F. E., W. L. THOMPSON, AND J.T. GRIFFITHS. 1949. Progress report on the fungus diseases of scale insects attacking citrus in Florida. The Florida Entomol. 32:1-11. FUXA, J.R., AND Y. TANADA. 1987. Epizootiology of insect diseases. Wiley Interscience Publ., NY. 555 p. GOULI, V.V., S.Y. GOULI, S.D. COSTA, AND M.V. SHTERNSHIS. 2005. Comparison of wash, leaf imprint and adhesive tape methods for estimation of fungal spore deposition on leaves. Mycol. and Phytopathol. 39 (3):99-103. HILLIS, D.M., AND M. T. DIXON. 1991. Ribosomal DNA: molecular evolution and phylogenetic inference. Q. Rev. Biol. 66:411–453. HUMBER, R.A. 1997. Fungi: identification. P: 153-185 in Lacey, L., editor. Manual of techniques in insect pathology. Academic Press. London. LATUNDE-DADA, A.O., 2001. Colletotrichum: tales of forcible entry, stealth, transient confinement and breakout. Mol. Plant Pathol. 2 (4):187-198.

LORD, J.C., 2005. From Metchnikoff to Monsanto and beyond: The path of microbial control. J. Invertebr. Pathol. 89:19-29. MCCLURE, M.S., AND M.B FERGIONE. 1977. Fiorinia externa and Tsugaspidiotus tsugae (Homoptera: Diaspididae): Distribution, abundance and new hosts of two destructive scale insects of eastern hemlock in Connecticut. Environ. Entomol. 6:807-811. MCCLURE, M.S. 1977. Parasitism of the scale insect, Fiorinia externa (Homoptera: Diaspididae), by Aspidiotiphagus citrinus (Hymenoptera: Eulophidae) in a hemlock forest: Density dependence. Environ. Entomol. 6:551-555. MCCLURE, M.S. 1986. Population dynamics of Japanese hemlock scales: A comparison of endemic and exotic Communities. Ecology 67 (5):1411-1421. MCCLURE, M.S. 2002. The elongate hemlock scale, Fiorinia externa Ferris (Homoptera: Diaspididae): a new look at an old nemesis. P: 248-253 in Proceedings of the hemlock woolly adelgid in the eastern United States symposium, Onken, B., R. Reardon, and J. Lashomb (eds.). New Jersey Agricultural Experimental Station, Rutgers University, New Brunswick, NJ. MCCOY, C.W. 1985. Citrus: current status of biological control in Florida. P. 481- 499 in Biological Control in Agricultural IPM Systems. Academic Press, London, NY. MCCOY, C.W., R.A. SAMSON, and D.G. BOUCIAS.1988. Entomogenous fungi. P. 151-236 in Handbook of natural pesticides, Vol. V. Microbial Pesticides. Part A. Entomogenous protozoa and fungi, Ignoffo, C. M., and N. B. Mandava (eds.), CRC Press, Boca Raton, FL. MILLER, J. H. 1940. The genus Myriangium in North America. Mycologia 32:587-600. NIKOH, N., AND T. FUKATSU. 2000. Interkingdom host jumping underground: phylogenetic analysis of entomoparasitic fungi of the genus Cordyceps. Mol. Biol. Evol. 17:629-638. O’DONNELL, K. 1992. Ribosomal DNA internal transcribed spacers are highly divergent in the phytopathogenic ascomycete Fusarium sambucinum (Gibberella pulicaris). Curr. Genet. 22:213–220. O’DONNELL, K. 1993. Fusarium and its near relatives. P. 225–236 in The fungal holomorph: mitotic, meiotic, and pleomorphic speciation in fungal systematics, Reynolds, D.R., and J.W. Taylor. (eds.). CAB International, Wallingford, UK. PARKER, B. L., M. SKINNER, V. GOULI, S. GOULI, J. MARCELINO, J. CARLSON AND L. SCHWARTZBERG. 2005. Management of elongate hemlock scale with entomopathogenic fungi. P. 161-168 in 3rd Symposium on Hemlock Woolly Adelgid in the Eastern United States, Onken B., and R. Reardon (eds.). USDA For. Ser. SAMSON, R.A., H.C. EVANS, AND J-P. LATGÉ. 1988. Atlas of entomopathogenic fungi. Springer-Verlag, Berlin. 187 p. SANDS, D.P.A., AND R.G. Van DRIESCHE. 2000. Evaluating host specificity of agents for biological control of arthropods: rationale, methodology and interpretation. P. 69– 83 in 10th International symposium on biological control of weeds, Van Driesche, R.G., T.A. Heard, A.S. McClay, and R. Reardon (eds.). For. Ser. Bul. FHTET-99-1, Morgantown, WV. SCHMITTHENNER, A.F., AND J.W. HILTY. 1962. A modified dilution technique for obtaining single-spore isolates of fungi from contaminated material. Phytopathol. 52:582–583.

TALHINHAS, P., S. SREENIVASAPRASAD, J. NEVES-MARTINS, AND H. OLIVEIRA. 2002. Genetic and molecular characterization of Colletotrichum acutatum causing anthracnose of lupins. Phytopathol. 92:86-996. TANADA, Y. AND H.K KAYA. 1993. Epizootiology. P. 595-632 in Insect pathology, Tanada Y., and H. K. Kaya (eds.). Academic Press, New York. USDA FOREST SERVICE. 2004. Forest insects and diseases conditions in the United States 2003. U.S. Depart. of Agric. For. Serv. Pub. 142 p. VAN EPENHUIJSEN, C.W., R.C. HENDERSON, A. CARPENTER, AND G.K. BURGE. 2000. The rise and fall of manuka blight scale: a review of the distribution of Eriococcus orariensis (Hemiptera: Eriococcidae) in New Zealand. N. Z. Entomol. (23):67-70. WATSON, J.R., AND E.W. BERGER.1937. Citrus insects and their control. University of Florida, Agric. Exp. Sta. Bul. 88. 135 p. WHITE, T.J., T. BRUNS, S. LEE, AND J.W. TAYLOR. 1990. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. P. 315-322 in PCR Protocols: A Guide to Methods and Applications, Innis M.A., D.H. Gelfand, J.J. Sninsky, and T.J. White (eds.). Academic Press, Inc., New York.

CHAPTER 3

Colletotrichum acutatum f. sp. fiorinia infection of a scale insect, Fiorinia externa Ferris (Hemiptera: Diaspididae)

José Marcelino1

Entomology Research Laboratory, University of Vermont, 661 Spear Street, Burlington, VT 05405

Rosanna Giordano

Illinois Natural History Survey, Division of Biodiversity and Ecological Entomology, 1816 S. Oak St.

Champaign IL, 61820

Svetlana Gouli

Entomology Research Laboratory, University of Vermont, 661 Spear Street, Burlington, VT 05405

Vladimir Gouli

Entomology Research Laboratory, University of Vermont, 661 Spear Street, Burlington, VT 05405

Bruce L. Parker

Entomology Research Laboratory, University of Vermont, 661 Spear Street, Burlington, VT 05405

Margaret Skinner

Entomology Research Laboratory, University of Vermont, 661 Spear Street, Burlington, VT 05405

David TeBeest

Department of Plant Pathology, Plant Science Building, University of Arkansas, Fayetteville,

Arkansas 72701

Roberto Cesnik

EMBRAPA - Embrapa Meio Ambiente, CP 69, CEP 13820-000, Jaguariúna-SP, Brazil

1 Email: [email protected]

76 Abstract: An epizootic has been reported in Fiorinia externa populations in New York,

Connecticut, Pennsylvania and New Jersey. Infected insects have profuse sclerotia

masses enclosing their bodies. The most commonly isolated microorganism from these

infected F. externa was Colletotrichum sp. A morphological and molecular

characterization of this fungus indicated that it is closely related to phytopathogenic C.

acutatum isolates. Isolates of Colletotrichum sp. from F. externa in areas of the epizootic were genetically similar and were named Colletotrichum acutatum f. sp. fiorinia based on our findings. In vitro and in planta matings observed between isolates of C. acutatum f. sp. fiorinia could serve as a possible source of genetic variation may give rise to new biotypes with a propensity to infect insects. Only one other strain, C. gloeosporioides f. sp. ortheziidae has been reported to be entomopathogenic.

Key Words: Fiorinia externa, Colletotrichum acutatum f. sp. fiorinia, entomopathogenic fungi, epizootic

Introduction

The ascomycete Colletotrichum Corda (1831) is a widely occurring and intensively studied plant pathogen and research has focused on its effects on plants

(Sutton, 1992). There is one reported case of a member of this genus, C. gloeosporioides, that causes epizootics on the scale Orthezia praelonga Douglas (1891) (Hemiptera:

Coccoidea), a major pest of citrus in Brazil (FIG. 3.1S). It was first identified by Batista

and Bezerra (1966) and morphologically confirmed by the Commonwealth Agriculture

Bureau International, CABI (H.C. Evans, CABI report 1990). Biological control of O.

praelonga, using C. gloeosporioides, has been conducted in Brazil (Viegas et al 1995,

77 Cesnik and Ferraz 2000) and scale mortality of 85-96% by 70 d after application has been

obtained (Cesnik et al 1996).

For most Colletotrichum spp., the complete host range is unknown (Fagbola and

Abang 2004). This genus is widely encountered in humid and sub-humid tropics (Mills et al 1992, Waller 1992). Reports of new hosts are common. In New Zealand 39 hosts from

23 plant families have been reported (Simmonds 1965). Xiao et al (2004) recovered this genus from 23 cultivated and non-cultivated hosts in west-central Florida. In Japan,

Moriwaki et al (2002) isolated 25 Colletotrichum spp. from 123 species including

Gramineae, Poaceae, Leguminosae and Cucurbitaceae. An extensive record of

Colletotrichum hosts was made by Lenné (1992), identifying nine species of the fungus associated with 102 species of plants (Sutton, 1992).

Colletotrichum spp. employ different infection strategies in plants ranging from endophytic to hemibiotrophic and subcuticular intramural growth (Smith et al 1999). In many of the reported hosts, quiescent mesophyll intercellular mycelia rather than necrotrophic growth is observed (Wei et al 2004).

This paper presents the second report of a member of the genus Colletotrichum infecting an insect and the first report of infection of an insect by a species within

Colletotrichum in North America. In 2002 an epizootic associated with a complex of fungi was observed in populations of elongate hemlock scale, Fiorinia externa Ferris

(Hemiptera: Coccoidea), a pest of eastern hemlock [Tsuga canadensis (L.) Carrière]

(McClure, 2002). The fungi associated with this epizootic appear to be of importance in the reduction of the F. externa population which together with the hemlock woolly

78 adelgid, Adelges tsugae Annand (Hemiptera: Adelgidae) are causing a significant decline in what has been one of the dominant components of New England’s forests for the last

8000 yr (Foster and Zebryk 1993, Danoff-Burg and Bird 2002, Snyder et al 2002).

Black sclerotia masses covering 80-100% of F. externa populations were

observed (FIG. 3.1T). Mortality of F. externa reached 36.8% (Marcelino et al. submitted).

Although the geographical origin of the epizootic is unknown presently it has been detected in 36 sites in New York, Connecticut, Pennsylvania and New Jersey.

The objective of this work was to morphologically, biologically and molecularly characterize the Colletotrichum sp. isolated from F. externa and to compare it to selected plant pathogenic strains in the same genus.

Materials and Methods

Morphological plasticity.—Morphological plasticity of 26 pure cultures of

Colletotrichum sp. isolated from F. externa collected in Bayberry lane; Mohonk; Esopus and Ward Pound Ridge Reservation, in New York state, grown on potato dextrose agar media (PDA) maintained at 10, 15, 20 and 22 C was determined. The following fungal

characteristics were visually monitored after 10 d: mycelium color, mycelium

pigmentation, media pigmentation and presence and abundance of conidia masses.

Monoconidial isolates and isolates derived from fungal suspensions in sterile distilled

water were inoculated into 20 ml Potato Dextrose Agar (PDA) medium and cultured in

Petri dishes. A stage-mounted DC3001 micromanipulator (World Precision Instruments,

Inc., Sarasota, Florida, US) was used to obtain monoconidial isolates. The experiment was repeated three times for all isolates.

79 Sexual recombination.—Cross-fertile sexual recombination was attempted in vitro by testing all combinations of crosses between the following 20 d old cultures:

Colletotrichum sp. isolates from the same and different localities of the epizootic; the single known entomopathogenic strain of C. gloeosporioides from Brazil (ARSEF4360); two plant pathogenic strains, ERL1395 and ERL1385, isolated from a golden delicious

apple and a tulip tree (Liriodendron tulipifera L.) growing in the area of the epizootic

(TABLE 3.1).

In vitro crosses were done on minimal salts medium (MSM), according to the

protocol by Guerber and Correll (2001). Petri dishes were sealed with parafilm. Plates

were incubated at room temperature (22 ± 0.5 C) with two constant cool white

fluorescent lights placed 1.2 m above the plates and four additional lights 2.5 m away on

the surrounding walls. Autoclaved birch wood (Betula sp.) toothpicks (Diamond Brand,

Inc., Minneapolis), and excised 2 cm long strawberry stems sterilized with propylene

oxide, were used as substrates to assess cross fertilization. Substrates were arranged in an

N pattern. Mating plates were screened after 40 d to detect the presence of perithecia in

the middle toothpick or stem, where the probability of selfing was reduced. The cross-

fertilization experiment using toothpicks was repeated twice, whereas the one with

strawberry stems was conducted once.

In planta sexual crossings were performed according to protocols from Cisar et al

(1996), using 4.5 yr old hemlock seedlings [Tsuga canadensis (L.) Carrière], three month

old bush snap bean (Phaseolus vulgaris L. var. Blue Lake 274) and three month old

strawberry plants (Fragaria x ananassa Duchesne var. Honeoye) to determine if

80 Colletotrichum sp. (EHS58) obtained from F. externa and Colletotrichum sp. (ERL1385), isolated from a tulip tree could self and/or cross-fertilize. Strains of both fungi used in these experiments originated from an area of high incidence of the epizootic (Ward

Pound Ridge Res., NY). Strain ERL1385 was used because this Colletotrichum sp. was

repeatedly isolated from tulip trees and because molecular data confirmed that it was

identical to the Colletotrichum strain EHS58 isolated from F. externa. Two parallel sets of

plants were set up. The first set included six plants per species while the second set

included four. Plants were individually bruised with a sterile scalpel (four bruises/plant)

6 and inoculated with 200 µl of 10 suspension of either Colletotrichum sp. EHS58 or

Colletotrichum sp. ERL1385. Inoculated plants were held in a dew chamber (3.95 m high ×

1.10 m wide × 1.10 m deep built with 2.5 cm diam PVC pipe and covered with plastic) for 24 h at 22 ± 1 C according to protocols of TeBeest (1988), and then placed in a

greenhouse at 22 C for 8 d. After this period, the same bruises were inoculated with the

alternate strain (ie EHS58 followed by ERL1385 or ERL1385 followed by EHS58 depending

on the first strain initially inoculated). Re-inoculated plants were again held in the dew

chamber for 24 h then moved to the greenhouse. For the first set of plants (six replicates),

sample stems were excised weekly, whereas for the second set of plants (four replicates)

the stems were excised at 3 and 6 wk. A 2 cm sample was excised from each plant and

placed in MSM medium with penicillin (5ml/l) and streptomycin (12.5ml/l), and

maintained in a growth chamber at a 8:16 h photoperiod (L:D) and 22 C. They were

examined for the presence of the teleomorph stage (Glomerella) after 1 wk. The

81 experiment above was repeated using the bean plants, and inoculated with ERL1385 and

another entomopathogenic strain, EHS48 from Bayberry Lane, NY.

Measurement of fungal structures.—Conidial spores were harvested from 14 d old

monoconidial PDA cultures of 26 Colletotrichum sp. from the epizootic in the US and two Colletotrichum f. sp. ortheziidae from the epizootic in Brazil. Spores were stained with cotton blue lactophenol to enhance contrast. Appressoria were produced on 1 cm dm strawberry leaf discs. A drop of culture suspension was placed on individual leaf discs.

Samples were incubated for 24 h at 22 C in Petri dishes lined with moist filter papers.

Appressoria were removed from leaves using the scotch tape print technique of Gouli et al (2005), stained with cotton blue lactophenol to enhance visualization and avoid plant tissue interference. Conidia and appressoria were viewed using a 40x phase-contrast

Trinocular BH2 Olympus compound microscope and photographed using a CCD digital camera (Pixelink, Vitana Corporation, Ottawa, Canada). Measurements of length and width were taken (Nconidia=1751; Nappressoria=1387), from which shape and elliptical form factors were calculated using the formulas below (Metamorph® software, Universal

Imaging Corp., West Chester, Pennsylvania). Intact perithecia were removed from bean

stems using a sterile insect pin and then placed in a glass slide, submerged in a droplet of

distilled water and mounted with a cover slide and photographed at 40x in an Olympus

BX51 photomicroscope connected to an AxioCam HC camera (Carl Zeiss Inc.,

Oberkochen, Germany). Ascospores extruded from perithecia were viewed in distilled

water and photographed using the same procedures. Length, width, area and shape were

calculated for both perithecia (N=40) and ascospores (N=119). In addition, the elliptical

82 form factor was calculated for the perithecia. The following values were calculated using

Metamorph® software:

Measurements:

1   Spore length =  P + P2 −16A  4  

1   Spore width =  P − P2 −16A  4  

Shape Factor = 4πA (Circumference =1) P2

Elliptical Form Factor = Length (Ratio of object’s breadth to its length) Breadth

P = Perimeter; A = Area

Appressorium length = Longest chord through the object

Appressorium width = Horizontal dimension of the object

Adapted from Metamorph® (2003)

Statistical analysis.— The SNK test (P=0.05) was used to identify whether there was a

significant difference between the morphological parameters obtained for the isolates

tested. Statistical analysis was performed using SAS® software (SAS Institute 1990).

Phylogenetic analyses.—Six Colletotrichum sp. pure strains obtained from F. externa

were chosen for molecular analysis based on data for germination, growth rate and

conidial production (Parker et al 2005). The rDNA from the following entomopathogenic

and phytopathogenic strains was also sequenced: C. gloeosporiodes. f. sp. ortheziidae

(ARSEF4360), from the USDA-ARS collection of entomopathogenic fungal cultures; C.

83 gloeosporiodes. f. sp. ortheziidae from EMBRAPA, Brazil (EMA26) and C. acutatum,

ERL1379 and ERL1380 (TABLE 3.1).

The following genes were used to characterize selected isolates: D1/D2 region of

the 28S ribosomal DNA, using primers NL1 and NL4 (O’Donnell 1992, 1993) commonly used for phylogenetic analysis at the genus level and above (Hillis and Dixon 1991); internal transcribed spacers ITS1 and ITS2, using primers ITS1 and ITS4 (White et al

1990) for within species differentiation (Afanador-Kafuri et al 2003); introns of the

glyceraldehyde-3-phosphate dehydrogenase (GPDH), associated with the infection

process of Colletotrichum (Wei et al 2004), with primers GDF1 and GDR1 (Templeton et

al 1992); glutamine synthetase protein (GS), highly expressed during pathogenesis using

primers GSF1 and GSR1 (Stephenson et al 1997), and newly constructed internal primers

GLUintF1 (5′ -AGCCGGAAGTCGGAGACATCCCG- 3′) and GLUintR2 (5′ -

CGTTGCTGTTCTCCACGCAAT- 3′). The β-Tubulin 2 protein encoding gene, used to

distinguish fungi at deep phylogenetic levels (Thon and Royse 1999, Lubbe et al 2004),

was amplified with primers TB5 and TB6 (Pannacione and Hanau 1990). The mating-

type gene (MAT 1-2) from the High Mobility Box (HMG), used to study fungal biology

(Turgeon 1998) was amplified with primers HMGacuF and HMGacuR (Du et al 2005).

A genetic characterization of representative Colletotrichum sp. isolates from the different regions of the epizootic, the Brazilian entomopathogenic C. gloeosporioides (ARSEF4360) and the phytopathogenic C. acutatum (ERL1379) was done using randomly amplified

polymorphic DNA (RAPDs), with primers derived from repeated sequences GACA(4)

84 (Weising et al 1989), GACAC(3) (Gupta and Filner 1991) and ACTG(4) (Freeman et al

1995).

DNA was extracted from 1 wk old cultures using the Power SoilTM DNA kit (Mo

Bio Laboratories, Inc., Carlsbad, California), and the FastPrepTM FP120 machine

(Thermo Savant, Holbrook, New York). Samples were shaken for 5 min at 5.5 m/s to break open fungal cell walls. The following modification was made to the Power SoilTM

DNA kit; DNA was eluted using 100 µl of diluted elution buffer AE from Qiagen (1:15) containing 10 mM of Tris and 0.5 mM of EDTA, and concentrated down to 20 µl with a

speed vacuum (Eppendorf Centrifuge 5415C, Vaudaux, Schönenbuch, Switzerland).

Polymerase Chain Reaction (PCR) was performed using Ready-To-Go PCR beads (Amersham Biosciences Inc., Piscataway, New Jersey). Genes were amplified

using the following protocol: initial denaturation at 95 C for 2 min followed by 30 cycles

of 95 C for 30 s (denaturation), 50 C for 30 s (annealing) and 72 C for 1 min (elongation)

with the following changes in annealing temperature: ITS (52 C); GPDH (54 C); β-

Tubulin 2 (65 C) and MAT 1-2 (55 C). RAPDs were amplified using Freeman et al.

(1995) protocols. PCR products were purified using the Qiagen QIAquick PCR purification kit (Valencia, CA) or Princeton Separations Centri SpinTM columns

(Adelphia, New Jersey). DNA was stored at 4 C. PCR products were sequenced using

BigDye v1 and BigDye v3 terminator cycle sequencing kit (Applied Biosystems, Foster

City, California) using the following protocol: initial denaturation at 95 C for 3 min

followed by 30 cycles of 95 C for 10 s (denaturation), 50 C for 5 s (annealing), and 60 C

for 2 min (elongation).

85 Sequencing reactions were completed on a 3130xl Genetic Analyzer (Applied

Biosystem, Foster City, California). Chromatograms were edited, and contiguous

sequences were generated, using SequencherTM (Gene Codes Corporation, Ann Arbor,

Michigan). Sequences were analyzed with related sequences obtained from GenBank

(Templeton et al 1992, Dufresne 1997, Nirenberg et al 2002, Lubbe et al 2004, Du et al

2005, Talhinhas et al 2002, 2005) with the exception of the majority of GPDH and GS

sequences which were not available in GenBank and were manually transcribed from Liu

(2002). Introns were aligned individually with the aid of ClustalW (Chenna et al 2003)

and retrieved with Jalview software (Clamp et al 2004. Exons were aligned separately

based on their respective protein sequences using McClade (Maddison & Maddison

1992). Since, rDNA analyses have not been sufficient, so far, to resolve phylogenetic

relationships in Colletotrichum spp. (Crouch et al 2005, Du et al 2005) the

Colletotrichum spp. used as outgroups were based on their genetic distance to the strains

used in the phylogenetic analysis. The following outgroup taxa were used: C. malvarum

(Z18981) for the D1/D2 domain of the 28S gene; C. malvarum (949C3E55) and C. gloeosporioides (9E1589A5) for the GS gene; C. kahawae (AY376588) and C.

gloeosporioides (AY376582); Glomerella cingulata (M93427) and G. cingulata

(7BDBDAF86) for the GPDH gene; C. coccodes (AY3766528) for the ITS region and C.

gloeosporioides (DQ002823) for the HMG at the MAT1-2 gene.

Phylogenetic trees were estimated using maximum parsimony as implemented in

PAUP 4.0b10 (Swofford, 2002). One representative isolate was included in the analysis

because the genomic sequences of Colletotrichum sp. from all the regions of the epizootic

86 were identical for all genes analyzed. Sequence gaps were treated as missing data.

Bootstrap analysis was performed using 1000 Bootstrap replications with 30 random

additions of taxa. Multiple equally parsimonious trees where combined into a single strict

consensus tree. Only bootstrap values above 70 were included. Sequences used in this

® analysis were deposited in GenBank (accession numbers included in TABLE 3.1). The

sequence alignment for all the genes sequenced and respective phylogenetic trees, were

also deposited in TreeBase databases (accession numbers to be assigned). Primers were

excluded from published sequences and sequence alignments.

Results

Morphological plasticity.—The isolates used in this study showed a wide range of

morphological variability often presenting several phenotypes as sectors within a colony

(FIGS. 3.1J-N), for both single-spore and suspension derived isolates (TABLE 3.2).

Mycelium color ranged from gray or black when cultured on PDA at 10 and 15 C

to pink or orange at 20 and 25 C. Multiple colors were at times present in the mycelia and

the media (FIG. 3.1M). Chromogenic media pigmentation, typical of C. acutatum, was

observed at 20 and 25 C in most of the isolates with gray and/or pink mycelium (FIG.

3.1O). Single spore isolates showed a stronger black color pigment in the media at 10 C

than the suspension-derived isolates, which produced a gray color at 10 C. Only isolates

EHS46, EHS50 and EHS51 produced orange mycelium consistently at all test temperatures.

For the remaining isolates, gray mycelium pigment was observed at all temperatures.

Aerial mycelium was seldom observed at all temperatures and isolates. Conidial masses

(FIG. 3.1P) were not observed on single-spore cultures but did occur in cultures derived

87 from suspension at 20 C (TABLE 3.2). Statistical analysis was not performed on these

data, as only the presence or absence of color pigment was recorded. In general, isolates

grown at 10 C or 15 C (both single-spore or suspension-derived) produced gray

mycelium, with sectors of other colors. In contrast, at 25 C, most of the isolates produced

an equal combination of gray and pink mycelia with local sectors of other colors. Single-

spore isolates were more uniform in color although color differences were common at 15

and 25 C.

Sexual recombination.—In vitro crossings were partially successful when using

toothpicks as a substratum (TABLE 3.3). Cross-fertile recombination on toothpicks

appeared to be incomplete in both repetitions of the experiment although tri-dimensional

sterile structures were observed together with profuse conidial masses (FIG. 3.1R)

especially on the diagonally-placed toothpicks (N). For EHS48 79% of the attempted

crosses resulted in the production of these sterile structures. The phytopathogenic isolate

(ERL1395) did not produce sterile structures in any of the crosses with the epizootic

strains. The tulip tree isolate (ERL1385) produced sterile structures in 64% of the

attempted crosses. The number of sterile structures was observed to be higher in crosses

with isolates from different geographic origins. The cross-fertile bioassay using

strawberry stems as a substratum produced profuse conidia masses (FIG. 3.1Q).

In planta cross-fertilizations with beans, strawberries and beans as substratum were successful only in beans. At 4 wk, and after 1 wk from excision from the plant, profuse numbers of perithecia were observed in self-fertile cross of EHS58 × EHS58. The cross-fertile ERL1385 × EHS58 produced perithecia after 5 wk in planta (FIG. 3.1A) and 1

88 wk after excised stems in petri-dishes (FIG. 3.1D). Perithecia were produced in planta, in two of the stem cuts in one of the experimental bean plants and in vitro in two bean stems cultured in MSM for 1 wk. Perithecia were not produced in a repeated experiment on beans. However, the perithecia produced in the bean stems in the first experiment were fertile, generating asci containing eight visible ascospores (FIG. 3.1B).

Measurement of fungal structures.— Wide ranges in the dimensions of the different

propagules were measured. The mean conidial length of the 26 entomopathogen

Colletotrichum sp. strains (FIG. 3.1I), was significantly smaller (5.61-8.57 × 2.73-4.22

µm) than the reported means for C. acutatum (TABLE 3.4). The ranges of means for

Brazilian isolates of C. gloeosporioides f. sp. ortheziidae, EMA26 (FIG. 3.1G) and

ARSEF4360 (FIG. 3.1H) were 10.35 × 2.50 µm and 11.88 × 2.37 µm, respectively. These

ranges were significantly lower than those obtained for the Colletotrichum sp. isolates

from sampled epizootic areas, with the exception of isolates EHS51, EHS52 and EHS56, which were within the range of widths for EMA26. Conidia of C. gloeosporioides f. sp.

ortheziidae (EHS35) were more oblong shaped than the entomopathogenic Colletotrichum

sp. from F. externa. In addition, the range of means for C. gloeosporioides f. sp.

ortheziidae was also smaller than that reported for C. gloeosporioides. Only one strain of

Colletotrichum sp., EHS41, was significantly different from all the other strains in terms of the spore area (TABLE 3.4).

The mean appressoria size (length × width) of the entomopathogenic

Colletotrichum sp. (FIG. 3.1E), ranged from 6.35-7.85 × 5.58-6.85 µm and was similar to

the reported range for C. acutatum (TABLE 3.5). The appressoria obtained from C.

89 gloeosporioides f. sp. ortheziidae EMA26 (FIG. 3.1F), ranged from 8.11 × 6.90 µm and

were also within the range for this species. C. gloeosporioides, ARSEF4360, did not produce appressoria after numerous attempts on strawberry leaves, poinsettia leaves,

Euphorbia pulcherrima and Citrus sp. leaves and fruit or mango, Mangifera sp., fruit).

Although we found significant differences among isolates in the appressoria’s shape, they

fell within the reported range in most cases. An average shape factor of 0.74-0.85 µm

was calculated for Colletotrichum sp. (spherical=1) and 0.71 in EMA36. The spore area for the 26 Colletotrichum strains and two C. gloeosporioides f. sp. ortheziidae strains were statistically identical (TABLE 3.5). Perithecia obtained from cross-fertile strains on

plant stems after 7 d on MSM, averaged 115.39 × 108.83 µm, which was not within the

reported range of Glomerella acutata. However, perithecia obtained from stems averaged

198.68 × 183.75 µm which is within the reported range for G. acutata. Perithecia

retrieved in planta and from stems in MSM media were statistically different for length

and width, shape and area (P<0.05) (TABLE 3.6).

No ovoid perithecia were found among specimens retrieved from plant stems,

whereas in Petri dish cultures 21% of 26 perithecia were ovoid. The ratio of perithecium breadth to length (elliptical factor) was not statistically different for perithecia in planta

and on MSM. The length was greater than the width, with a more marked difference in

the perithecia from MSM. The area of the perithecia produced in planta (32211.60 µm2) was significantly different from those produced on MSM (8639.28 µm2). Ascospores

discharged from all perithecia measured were smaller than those reported for G. acutata

(TABLE 3.6). However, differences in area were found between ascospores from

90 2 perithecia in planta (FIG. 3.1C), 28.30 µm and those produced on stems and cultured on

2 MSM (39.29 µm ). These differences were also found in the shape factor (TABLE 3.7).

Molecular analysis.—A RAPDs analysis of seven representative isolates of

Colletotrichum sp. from the epizootic and a single phytopathogenic C. acutatum isolated from blueberry (ERL1379) showed identical band patterns (FIG. 3.2). However, some

genetic variation was observed, between the strains of Colletotrichum from the epizootic area in the northeastern US and the Colletotrichum sp. from the entomopathogenic C.

gloeosporioides collected in Brazil (ARSEF4360).

We sequenced six genes from seven strains of Colletotrichum isolated from the F. externa epizootic and two C. gloeosporioides f. sp. ortheziidae strains from the epizootic in Brazil. These genes comprise a total of 3121 base pairs. Parsimony informative characters for the respective genes used in the analysis were as follows: GPDH (72.1%);

HMG at the MAT1-2 (56.6%); GS (34.6%); β-Tubulin (26.2%); ITS (16.8%) and D1/D2 region of the 28S rDNA (3.4%). Colletotrichum and C. gloeosporioides nucleotide sequences obtained for the six genes where individually compared with related sequences from GenBank using BLAST. A subset of sequences with similarity at or above 90% were retrieved from GenBank and incorporated in the data set used for the phylogenetic analysis included herein. Colletotrichum isolated from F. externa and C. gloeosporioides isolated from O. praelonga were found to be most similar to known representative phytopathogenic C. acutatum species. For the ITS sequences, Colletotrichum from F. externa was identical to C. lupini (FIG. 3.3). The mean character difference for the

respective genes used in the analysis between the two entomopathogenic Colletotrichum

91 spp., listed in decreasing order, is as follows: GPDH (8.4%); GS (5.9%); β-Tubulin2

(4.1%); HMG at MAT1-1 (3.7%); ITS (1.3%) and the D1/D2 region of the 28S rDNA

(0.36%). We could not analyze a concatenated data set of all of the genes because sequences are not available for the same taxa in all genes. With the exception of the two ribosomal sequences, ITS and the 28S, the analysis from all other genes analyzed show strong support for the placement of the two entomopathogenic forms within a monophyletic C. acutatum, despite the different taxa used for each gene (FIG. 3.4, 3.5 and

3.6). The GPDH gene showed the greatest level of divergence between the F. externa

derived Colletotrichum and C. gloeosporioides from O. praelonga, despite having sequenced 248 base pairs (FIG. 3.5). GPDH has the largest number of well supported

branches, whereas 28S had the least.

Discussion

The work herein reports on the occurrence of Colletotrichum acutatum f. sp.

fiorinia isolated from F. externa in 36 localities within the states of New York,

Pennsylvania, Connecticut and New Jersey. Twenty-six entomopathogenic isolates of

Colletotrichum sp. obtained from F. externa, originating from four different geographic

localities of the epizootic were morphologically characterized to determine whether they

were the same strain, and to compare them to known Colletotrichum species in order to

assess their closest relative. Included in this analysis is the only other known

Colletotrichum sp. to cause an epizootic in insects, C. gloeosporioides f. sp. ortheziidae,

obtained from two distant geographic localities from its epizootic areas in Rio de Janeiro

and São Paulo, Brazil.

92 Our morphological and molecular data indicated that the fungus isolated from F.

externa in the epizootic belonged to the genus Colletotrichum and that it was identical to the species C. acutatum. Colletotrichum affects a wide variety of plants (Lenné 1992,

Lardner et al 1999, Moriwaki et al 2002) with C. acutatum and C. gloeosporioides known to be the most cosmopolitan species. Reports of plant pathogens infecting insects are uncommon. Within the genus Colletotrichum there is only one other published case, that of C. gloeosporioides f. sp. ortheziidae infecting the scale Orthezia praelonga in Brazil

(Cesnik and Ferraz, 2000). Our data indicate that C. acutatum f. sp. fiorinia is associated with the epizootic in F. externa. The phylogenetic analysis obtained from four of the six genes, ITS (FIG. 3.3), GS (FIG. 3.4), GPDH (FIG. 3.5) and β-tubulin2 suggests that the

divergence in host utilization, from plant to insect, of both C. acutatum f. sp. fiorinia and

C. gloeosporiodies f. sp. ortheziidae are independent events. However, when using the

HMG at the MAT1-2 gene both (FIG. 3.6) taxa form a monophyletic group, perhaps due

to sampling error. Both Colletotrichum strains retrieved from F. externa and O.

praelonga appear to be derived from C. acutatum.

With the exception of conidia and ascospore size, there is congruence between the

two reliable morphological measurements and results obtained using molecular data.

Thus we name this fungus isolated from F. externa in the northeastern epizootic region of

the US is a C. acutatum f. sp. fiorinia.

We observed an array of phenotypes and sectors in cultures for Colletotrichum sp.

recovered from F. externa. The highest level of variability in pigmentation occurred at 20

C and the lowest at 10 C. Although fungal phenotypic plasticity is a common

93 phenomenon in vitro, this strain was unusual in that we observed up to five different

color pigments in both the mycelium and medium, in pure cultures. This level of

variation indicates a high rate of morphological heterogeneity in this strain. In contrast,

the two C. gloeosporioides f. sp. ortheziidae Brazilian strains tested did not display this

plasticity in culture, presenting a consistent orange (ARSEF4360) and gray mycelia

(EMA26) at both 15 C and 25 C.

Conidia of the 26 isolates of the entomopathogenic Colletotrichum sp., were

smaller (5.61 to 8.57 µm) than the reported length for the C. acutatum species complex.

The two entomopathogenic strains of C. gloeosporioides f. sp. ortheziidae, also produced

smaller conidia, than the reported for C. gloeosporioides species complex. Appressoria

length and width of both Colletotrichum sp. and C. gloeosporioides f. sp. ortheziidae

were consistently within the lower range of that reported for C. acutatum and C. gloeosporiodes. In both species appressoria were mainly ovoid.

Segregation of species based on morphological characters in the genus

Colletotrichum has been based primarily on measurements of appressoria. Although we found that conidia size for both entomopathogenic strains from the US and Brazil are outside the reported range for other Colletotrichum strains, we do not believe this result alone is a reliable means at differentiating species segregation because the standard method for measuring spores is inconsistent. The method involves placing propagules, suspended in water, on a slide, where they can be oriented in different directions or can move due to brownian motion, hence biasing readings. In contrast, these problems are avoided when measuring appressoria using the scotch-tape print technique (Gouli et al

94 2005). Therefore it is not surprising that the appressoria measurements made for

Colletotrichum sp. and C. gloeosporioides f. sp. ortheziidae fit within the range of

Colletotrichum albeit in the lower range of the spectrum.

We detected sexual reproduction in planta using snap bean as a substratum, in crosses between Colletotrichum sp. strains from F. externa and a tulip tree from the epizootic area. Successful sexual reproduction was also detected in vitro after one week culturing of stems excised from beans, and cultured on MSM. However, reproduction was stalled when birch toothpicks were used as a substratum and no reproduction was observed with strawberry stems. Selfing was detected in a self-cross with the entomopathogenic Colletotrichum sp. (strain EHS58 x EHS58). Our data cannot differentiate as to whether the sexual cross obtained between the Colletotrichum sp. strain from F. externa and the Colletotrichum sp. from a tulip tree was homothallic or heterothallic since we obtained the perfect stage (Glomerella) with different and same strain crosses. However, these crosses do indicate that sexual reproduction can occur in this strain, possibly resulting in the adaptation to new hosts (Guerber and Correll 2001) and generation of new biotypes.

The crosses observed in beans were fertile. Perithecia, produced by the perfect stage of Colletotrichum (= Glomerella) generated asci containing eight ascospores. The mean range for the perithecia (length × width) retrieved after excised stems where placed in MSM were not within the reported range of Glomerella acutata (115.39 × 108.83 µm), whereas perithecia retrieved directly from stems in planta were within the reported range of G. acutata (198.68 × 183.43 µm). Size differences were also observed with perithecia

95 retrieved from excised stems after one week of culturing on MSM, presenting a mean area almost four times smaller than the area of the ones retrieved directly in planta

(TABLE 3.6). Differences in the size of the ascospores produced in asci by the two types of perithecia were also found, with ascospores produced in perithecia directly retrieved from the stems being slightly smaller than the ones from perithecia in snap bean stems cultured in MSM. Incongruence between the size of perithecia and ascospores might result from differences in the environments in which the fruiting bodies were obtained.

Perithecia values within the reported range were obtained from plants reared in a greenhouse with natural photoperiod, whereas smaller and out of range values where obtained from plant material in Petri dishes, in MSM media, in a growing chamber at a constant 8:16 h photoperiod (L:D) and sealed with parafilm. Like the previously reported conidial spore measurements, and for similar reasons discussed above, the ascospores of

Glomerella sp. were not within the reported range for G. acutata.

Our molecular data indicate that a single population lineage of Colletotrichum from F. externa, henceforth referred as Colletotrichum acutatum f. sp. fiorinia, is present in the sampled epizootic area. A RAPDs analysis (FIG. 3.2) shows molecular homogeneity in C. acutatum f. sp. fiorinia strains sampled within the epizootic and a C. acutatum obtained from blueberry. However, differences were found between these apparently homologous strains and that obtained from C. gloeosporioides f. sp. ortheziidae.

A similar pattern was seen when six nuclear genes (ie the D1/D2 region of the

28S ribosomal DNA, ITS region, β-tubulin 2 gene, GPDH gene, GS gene and the High

96 Mobility Group of the mating-type gene, MAT1-2) were analyzed for these taxa. No differences were found between the C. acutatum f. sp. fiorinia strains collected within the

area of the epizootic and C. acutatum from a blueberry. As with the RAPDs analysis,

differences where found in the genes analyzed between C. acutatum f. sp. fiorinia and C.

acutatum strains and C. gloeosporioides f. sp. ortheziidae.

A BLAST search of data in GenBank using sequences of the nuclear genes listed

above, obtained from both C. acutatum f. sp. fiorinia and C. gloeosporioides f. sp.

ortheziidae, retrieved records of C. acutatum, some of which were identical. In addition

when using the ITS sequence we also retrieved another identical taxa, C. lupini

(AJ301968).

Data collected indicates that C. gloesporioides f. sp. ortheziidae appears to have

attained specificity for Orthezia praelonga (Teixeira et al 2001, Jonsson and Genthner

1997, Castro et al 1998). This special form is currently being effectively used as a

biological control agent in Brazil (Cesnik and Ferraz 2000) and under commercialization

(R. Cesnik pers comm). It is possible that these two strains of Colletotrichum are at

different stages of a host adaptation process. Data collected suggests that while C.

acutatum f. sp. fiorinia still retains some capacity to invade plants endophytically

(Marcelino et al. submitted), C. gloeosporioides f. sp. ortheziidae appears to have lost

this capacity.

It has been suggested that C. acutatum may have a broader host range than what

has been reported (Peres et al 2005). The cosmopolitan preference of this species for

plants may pre-adapt it to infect radically different hosts. The means by which such wide

97 range of preference in hosts can be achieved by this pathogen are uncertain (Wei et al

2004). The ability to expand host range can result from genetic variation subsequent to

sexual crossing. While we do not know whether this strain can produce an heterothallic

cross we have induced an homothallic cross and generated the sexual stage, Glomerella,

in our crosses. It is possible that this C. acutatum f. sp. fiorinia with its new propensity to

infect insects rather then plants may have resulted from same sex mating. This type of

reproductive strategy would produce meiotic clones, perhaps explaining the molecular

homogeneity of the C. acutatum f. sp. fiorinia strains sampled within the area of the

epizootic. A similar case of recombination via same-sex mating and subsequent expansion to new geographical niches has recently been reported for Cryptococcus gattii

(Fraser et al 2005).

In most Colletotrichum spp. affecting plants the prevalent mode of reproduction is

clonal, however, heterothallic intercompatibitiliy has been reported (Roca et al 2004,

Vaillancourt et al 2000, Crouch et al 2005).

The northeastern US forest area where the epizootic has occurred provides

optimal conditions for the growth of C. acutatum f. sp. fiorinia which may have

facilitated sexual crossing in this strain. Typical hemlock stands grow in riparian areas

that help create a microclimate where relative humidity as a rule can reach 80% in the

summer and attain 80 d/y of mist (Baldwin 1973, McGuire and Forman 1983).

The capacity of Colletotrichum acutatum f. sp. fiorinia to infect members of two

kingdoms widens the host range for this species. The classification of members of this

genus is in part determined by the identity of their hosts. The identification of a strain

98 which affects a wide variety of plants as well as an insect illustrates the difficulties inherent in using host identity for species segregation. This work indicates that multiple characters, including morphological and molecular, be used to classify member of this genus.

Acknowledgments

We thank Mr. Fred Little and Mr. Jeff Grosse for transcribing non-published genetic sequences in Liu, 2002. We also thank Dr. Felipe Soto for assistance with molecular analysis. This work was funded in part through a grant awarded by the Northeastern Area

State and Private Forestry, USDA Forest Service (# 04-CA-11244225286) and is in partial fulfillment of requirements for the PhD degree of J.M. at the University of

Vermont.

99 Literature Cited

Afanador-Kafuri L, Minz D, Maymon M, Freeman S. 2003. Characterization of Colletotrichum isolates from tamarillo, passiflora, and mango in Colombia and identification of a unique species from the genus. Phytopathology 93:579-587. Baldwin JL. 1973. Climates of the United States. U. S. Dep. Commerce, Washington, D. C. 113 p. Batista AC, Bezerra JL. 1966. Sobre o parasitismo de Colletotrichum gloeosporioides e outros fungos em Orthezia praelonga Douglas. Brotéria 35:68-74. (In Portuguese) Castro VL, Cesnik R and Oliveira, RCAL. 1998. Testes toxicológicos em ratos expostos ao Colletotrichum gloeosporioides. 6 Simpósio de Controle Biológico. Rio de Janeiro, Brazil. P 156. (In Portuguese) Cesnik R, Ferraz, JNG, Oliveira RCAL, Arellano F, Maia AH. 1996. Controle de Orthezia praelonga com o fungo Colletotrichum gloeosporioides isolado orthezia, na regiao de Limeira, SP. In: 5o Simpósio de Controle Biológico. Foz de Iguaçu, Brazil. P 363. (In Portuguese) ——,——. 2000. Orthezia praelonga, 1891 (Hemiptera, Ortheziidae) biologia, controle químico e biológico. Jaguariúna: Embrapa Meio Ambiente, Boletim de Pesquisa 9: 27 p. (In Portuguese) Cisar CR, Thornton AB, TeBeest DO. 1996. Isolates of Colletotrichum gloeosporioides (Telemorph: Glomerella cingulata) with different host specificities mate on Northern Jointvetch. Biol Control 7:75-83. Chenna R., Sugawara H., Koike T., Lopez R., Gibson TJ, Higgins DG., Thompson JD. 2003. Multiple sequence alignment with the Clustal series of programs. Nucleic Acids Res 31 (13):3497-500. Clamp M, Cuff J, Searle SM, Barton GJ. 2004. The Jalview java alignment editor. Bioinformatics 20:426-427. Corda AKJ. 1831. Die Pilze Deutschlands. In: Deutschlands Flora, 3. Abtheilung. Sturm, J. eds. 3 (12):33-64. Crouch JA, Clarke BB, Hillman BI. 2005. Diversity in natural populations of Colletotrichum from the tallgrass prairie. 23rd Fungal Genetics Conference. 15 to 20 March, 2005. Asilomar, California. (Abstract) Danoff-Burg JA, Bird S. 2002. Hemlock woolly adelgid and elongate hemlock scale: Partners in crime? In: Onken B, Reardon R, Lashomb J, eds. Proceedings of the Hemlock Wooly Adelgid in the Eastern United States Symposium, February 5-7, 2002; New Jersey Agricultural Experiment Station, Rutgers University, New Brunswick, NJ. p 254-268. Du M, Schardl CL, Nuckles EM, Vaillancourt LJ. 2005. Using mating-type gene sequences for improved phylogenetic resolution of Colletotrichum species complexes. Mycologia 97:641-658. Foster DR, Zebryk TM. 1993. Long-term vegetation dynamics and disturbance history of a Tsuga-dominated forest in New England. Ecology 74 (4):982-998. Fraser JA, Giles SS, Wenink EC, Geunes-Boyer SG, Wright JR, Diezmann S, Allen A, Stajich JE, Dietrich FS, Perfect JR, Heitman J. 2005. Same-sex mating and the

100 origin of the Vancouver Island Cryptococcus gattii outbreak. Nature 437:1360– 1364. Freeman S, Shabi E, Katan T. 1995. Characterization of Colletotrichum acutatum causing anthracnose of anemone (Anemone coronaria L.). App Env Microb 66 (12):5267- 5272. Gouli VV, Gouli SY, Costa SD, Shternshi MV. 2005. Comparison of wash, leaf imprint and adhesive tape methods for estimation of fungal spore deposition on leaves. J. Mycol. & Phytopath. 39:99-103. (In German) Guerber JC, Correll JC. 2001. Morphological description of Glomerella acutata, the teleomorph of Colletotrichum acutatum. Mycologia 93:216-229. Griffin MS, Sutheereland JR, Dennis JJ. 1987. Blight of conifer seedlings caused by Colletotrichum gloeosporioides. New Forest 1:81-88. Gupta M, Filner P. 1991. Microsatellites amplify highly polymorphic DNA bands in SPAR of plant DNA. In: Proc. Int. Soc. Plant Mol. Biol., Tucson, AZ. Abstr. 1705 Hillis DM, Dixon MT. 1991. Ribosomal DNA: molecular evolution and phylogenetic inference. Q Rev Biol 66:411–453. Hopkins JC, Lock W, Funk A. 1985. Colletotrichum acutatum, a new pathogen on western hemlock seedlings in British Columbia. Can Plant Dis Surv 65:11-13. Jonsson CM and Genthner FJ. 1997. Avaliação do potencial de patogenicidade e toxicidade do fungo entomopatógeno Colletotrichum gloeosporioides isolados de Orthezia em duas espécies de crustáceos. Bol. Pesqui. Embrapa-CNPMA 1:1-27. (In Portuguese) Lardner R, Johnston PR, Plummer KM, Pearson MN. 1999. Morphological and molecular analysis of Colletotrichum acutatum sensu lato. Mycol Res 103:275-285. Lenné JM. 1992. Colletotrichum diseases of legumes. In: Bailey JA, Jeger MJ, eds. Colletotrichum: Biology, Pathology and Control. Wallingford, UK: CAB International. p 135-166. Liu B. 2002. Molecular characterization and inter and intra species phylogenetic relationships in the fungal plant pathogen Colletotrichum. [Doctoral dissertation]. Fayetteville, Arkansas: Univ. Arkansas. 286 p. Lubbe CM, Denman S, Cannon PF, Groenewald JZ, Lamprecht SC, Crous PW. 2004. Characterization of species associated with diseases of Proteaceae. Mycologia 96 (6):1268-1279. Maddison WP, Maddison JR. 1992. MacClade: Analysis of phylogeny and character Evolution. Sinauer, Sunderland, MA. McClure MS. 2002. The elongate hemlock scale, Fiorinia externa Ferris (Hemiptera: Diaspididae): A new look at an old nemesis. In: Onken B, Reardon R, Lashomb J, eds. Proceedings of hemlock woolly adelgid in the eastern United States Symposium, February 5-7, 2002; New Jersey Agricultural Experiment Station, Rutgers University, New Brunswick, NJ. p 248-253. McGuire DA, Forman RTT. 1983. Herb cover effects on tree seedling patterns in a mature hemlock-hardwood forest. Ecology, 64:1367-1380.

101 Metamorph Imaging System (2003). Meta imaging series MetaMorph basic commands: Version 6.1 for Microsoft Windows 2000/XP: User's Guide. Universal imaging corporation, Downingtown, PA 19355. Mills PR, Sreenivasaprasad S, Brown AE. 1992. Detection and differentiation of Colletotrichum gloeosporioides isolates using PCR. FEMS Microbiol Lett 98:137- 144. Moriwaki J, Tsukiboshi T, Sato T. 2002. Grouping of Colletotrichum species in Japan based on rDNA sequences. J Gen Plant Pathol 68:307-320. Negasi AN, Parker BL, Brownbridge M. 1998. Screening and bioassay of entomopathogenic fungi for the control of silverleaf whitefly. Insect Sci Applic 18: 37-44. Nirenberg HI, Feiler U, Hagedorn G. 2002. Description of Colletotrichum lupini comb. nov. in modern terms. Mycologia 94: 307-320. O’Donnell K. 1992. Ribosomal DNA internal transcribed spacers are highly divergent in the phytopathogenic ascomycete Fusarium sambucinum (Gibberella pulicaris). Curr Genet 22:213–220. ——. 1993. Fusarium and its near relatives. In: Reynolds R., Taylor J. W., eds. The fungal holomorph: mitotic, meiotic and pleomorphic speciation in fungal systematics. Wallingford, United Kingdom. CABI International. p 225-233. Pannacione DG, Hanau RM. 1990. Characterization of two divergent β-tubulin genes from Colletotrichum graminicola. Gene 86:163-170. Parker BL, Skinner M, Gouli V, Gouli S, Marcelino JAP, Carlson J, Schwartzberg L. 2005. Management of elongate hemlock scale with entomopathogenic fungi. In: Onken B, Reardon, R, eds. 3rd Symposium on Hemlock Woolly Adelgid in the Eastern Unites States. Asheville, North Carolina. February 1-3, 2005. p 161-168. Peres NA, Timmer LW, Adaskaveg JE, Correll JC. 2005. Lifestyles of Colletotrichum acutatum. Plant Dis 89:784-796. Roca MG, Davide LC, Davide LMC, Mendes-Costa MC, Schwan RF, Wheals AE. 2004 Conidial anastomosis fusions between Colletotrichum spp. Mycol Res 108 (11): 1320-1326. SAS Institute. 1990. SAS/STAT user's guide. 4th ed. Cary, NC. SAS Institute. Sharon A, Yamaguchi K, Christiansen S, Horwitz BA, Yoder OC, Turgeon BG. 1996. An asexual fungus has the potential for sexual development. Mol Gen Genet 251:60-68. Simmonds JH. 1965. A study of the species of Colletotrichum causing ripe fruit rots in Queensland. Queensl J Agric Anim Sci 22: 437-459. Smith JE, Korsten LK, Aveling TASA. 1999. Infection process of Colletotrichum demantium on cowpea stems. Mycol Res 103 (2):230-234. Snyder CD, Young JA, Lemarie DP, Smith DR. 2002. Influence of eastern hemlock (Tsuga canadensis) forests on aquatic invertebrate assemblages in headwater streams.Can. J. Fish. Aquat. Sci. 59(2):262-275. Stephenson SA, Green JR, Manners JM, Maclean DJ. 1997. Cloning and characterization of glutamine synthetase from Colletotrichum gloeosporioides and demonstration of elevated expression during pathogenesis on Stylosanthes guianensis. Curr Genet 31:447-54.

102 Sutton, BC. 1992. The genus Glomerella and its anamorph Colletotrichum. In Bailey JA, Jeger MJ, eds. Colletotrichum – Biology, Pathology and Control. CAB International. Wallingford, UK. p. 1-26. Swofford DL. 2002. PAUP*. Phylogenetic analysis using parsimony (*and other methods). Version 10. Sinauer Associates, Sunderland, Massachusetts. Talhinhas P, Sreenivasaprasad S, Neves-Martins J, Oliveira H. 2002. Genetic and molecular characterization of Colletotrichum acutatum causing anthracnose of lupins. Phytopathology 92:986-996. ——, ——, Neves-Martins J, Oliveira H. 2005. Molecular and phenotypic analysis reveals the association of diverse Colletotrichum acutatum groups and a low level of C. gloeosporioides with olive anthracnose. Appl Environ Microbiol 71 (6):2987- 2998. TeBeest DO. 1988. Additions to host range of Colletotrichum gloeosporioides f. sp. aeschynomene. Plant Disease 72:16-21. Templeton MD, Rikkerink EH, Solon SL, Crowhurst RN. 1992. Cloning and molecular characterization of the glyceraldehyde-3-phosphate dehydrogenase-encoding gene and cDNA from the plant pathogenic fungus Glomerella cingulata. Gene 122(1): 225-230. Teixeira MA, Bettiol W and Cesnik R. 2001. Patogenicity of Coletotrichum gloeosporioides, Orthezia praelonga pathogenic agent, to citrus leaves, flowers and fruits. Summa Phytopathologica 27:352-357. (In Portuguese) Thon MR, Royse DJ. 1999. Partial β-Tubulin gene sequences for evolutionary studies in the Basidiomycota. Mycologia 91 (3):468-474. Turgeon BG.1998. Application of mating-type gene technology to problems in fungal biology. Ann Rev Phytopathol 36:115-137. Vaillancourt LJ, Du M, Wang J, Rollins J, Hanau RM. 2000. Genetic analysis of cross fertility between two self-sterile strains of Glomerella graminicola. Mycologia 92:430-435. Viegas EC, Sampaio HN, Carvalho POL, Perruso JC, Cassino PCR. 1995. Contole alternative de Orthezia praelonga Douglas, 1891 (Hemiptera, Ortheziidae) em laboratório. In: 15o Congresso de Entomologia, 12 a 17 Março 1995, Caxambú, Brazil. Pp: 333. (In Portuguese) Waller JM. 1992. Colletotrichum diseases of perennial and other cash crops: Bailey JA, Jeger MJ, eds. Colletotrichum – Biology, Pathology and Control. CAB International. Wallingford, UK. p 167-201. Wei YD, Shen W, Dauk M, Wang F, Selvaraj G, Zou J. 2004. Targeted-gene disruption of glycerol-3-phosphate dehydrogenase in Colletotrichum gloeosporioides reveals evidence that glycerol is a transferred nutrient from host plant to fungal pathogen. J Biol Chem 279:429-435. Weising K, Weigand F, Driesel AJ, Kahl G, Zischer H. 1989. Polymorphic simple GATA/GACA repeats in plant genomes. Nucleic Acids Res 17:10128. White TJ, Bruns T, Lee S, Taylor JW. 1990. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In: Innis MA, Gelfand DH,

103 Sninsky JJ, White TJ, eds. PCR Protocols: A Guide to Methods and Applications, Academic Press, Inc., New York. p 315-322. Xiao CL, MacKenzie SJ, Legard DE. 2004. Genetic and pathogenic analysis of Colletotrichum gloeosporioides isolates from strawberry and non-cultivated hosts. Phytopathology 94:446-453.

104

FIG. 3.1. Perithecia in planta from cross fertile strains of Colletotrichum sp., ERL1385xEHS58 (A); intact ascus (B); free ascospores (C); Perithecia discharging ascospores (D); appressoria of Colletotrichum sp., strain EHS47, from epizootic site (E); appressorium of C. gloeosporioides, strain EMA26, from Brazilian epizootic (F); Conidia from C. gloeosporioides, EMA26 (G) and ARSEF4630 (H); Colletotrichum sp., strain EHS48, from epizootic area (I); morphological plasticity and overlapping phenotypes of Colletotrichum sp. strains from epizootic localities (J-M); Top

(N) and reverse (O) of Colletotrichum sp. strain EHS58 in PDA at 25 C; Conidia produced by Colletotrichum sp. from epizootic site (P); Strawberry stem from cross fertile bioassay presenting profuse conidia masses (Q); Birch toothpick from cross fertile bioassay with conidia masses and sterile tri-dimensional structures (R); Adult O. praelonga infected with C. gloeosporioides f. sp. ortheziidae (orange masses), Brazil (S); Adult F. externa with epizootic, US (T). Bars: A = 200µm; B, C, E, F = 20µm; D = 60µm; G-I = 15µm, T = 5 mm.

105

FIG. 3.2. Random Amplified Polymorphic DNA (RAPDs) for Colletotrichum sp. strains from different

geographic localities within the epizootic area. EHS36 (A); EHS41 (B); EHS48 (C); EHS51 (D); EHS52 (E);

EHS58 (F); EHS61 (G); from a phytopathogenic C. acutatum from blueberry, ERL1379 (H); and from the

Brazilian entomopathogenic C.gloeosporioides f. sp. ortheziidae ARSEF4360 (I).

106 Colletotrichum sp. (F. externa) EF464580 C. lupini AJ301968

C. acutatum AY376519

C. acutatum AY376500 100 C. acutatum AY376497

Colletotrichum gloeosporioides (O. praelonga) EF593338 C. acutatum AF090853

C. acutatum AY376508

C. orbiculare AY376541 100 C. trifolii AJ301941

C. lindemunthianum AJ301958

75 C. caudatum AY376527 C. linicola AB046609

87 C. kahawae AY376540 100 C. gloeosporioides AJ311882

71 C. crassipes AY376530 C. boninense AY376569 79 100 C. capsici AY376526 C. truncatum AF451899

C. sublineolum AY376542 C. coccodes AY376528

FIG. 3.3. Phylogenetic tree (Inferred MP tree) among Colletotrichum spp. and Colletotrichum sp. strains isolated from Fiorinia externa, for the ITS region. Consensus of three equally parsimonious trees. Numbers above nodes are bootstrap values based on 1000 iterations with 30 random additions each. Only branches with values above 70 are considered well supported.

107 Colletotrichum sp. (F. externa) EF464580 98 C. acutatum 547012A1 C. acutatum 11F67OAF 84 C. acutatum FO6A7CC C. acutatum 9C2E3C5C

95 C. acutatum 14366523 C. gloeosporioides (O. praelonga) EF593338 100 C. acutatum 8AB8A03C C. acutatum 13A4BEAD C. acutatum E3D499S5 100 C. acutatum C464CEFA C. acutatum FA296838 C. acutatum 5863DBA1 89 C. acutatum D82818F7 C. acutatum E4F3130 C. acutatum B18896DA5 84 C. acutatum 3D2D89A9 C. acutatum 984DF794 100 98 95 C. acutatum PJ16-57BEA7F6 99 C. acutatum PJ11-57BEA7F6 99 C acutatum FD30092 C. acutatum J1ES33A3B 96 90 C. acutatum 247BF1FC7 C. acutatum 5F2CD5CE C. gloeosporioides 9E1589A5 C. malvarum 949C3E55

FIG. 3.4. Phylogenetic tree (Inferred MP tree) among Colletotrichum spp. and Colletotrichum sp. strains isolated from Fiorinia externa, for the GS gene. Consensus of nine equally parsimonious trees. Numbers above nodes are bootstrap values based on 1000 iterations with 30 random additions each. Only branches with values above 70 are considered well supported.

108 Colletotrichum sp. (F. externa) EF464580 96 C. acutatum DB5932DD

90 C. acutatum 17A1B49 99 C. gloeosporioides (O. praelonga) EF593338 94 C. acutatum 269A60EC 100 C. acutatum 3404C60C C. acutatum 63FF9E1A 73 95 C. acutatum B10CCE11 C. acutatum 11803ADD

100 C. sublineolum BrazilD47FE74 C. sublineolum SS1 Ac5CB1E2 100 C. orbiculare DQ792868 1

72 C. malvarum DQ792860 1 C. trifolii DQ792854 1 70 C. orbiculare DQ792866 1 100 C. malvarum DQ792870 1 C. orbiculare DQ792861 1 C. gloeosporiodes CA9B40F3 C. gloeosporiodes 9E1589A5

88 Glomerella cingulata M93427 Glomerella cingulata 7BDBDAF8 6

FIG. 3.5. Phylogenetic tree (Inferred MP tree) among Colletotrichum spp. and Colletotrichum sp. strains isolated from Fiorinia externa, for the GPDH gene. Consensus of 350 equally parsimonious trees. Numbers above nodes are bootstrap values based on 1000 iterations with 30 random additions each. Only branches with values above 70 are considered well supported.

109 Colletotrichum sp. (F. externa) EF464580

92 Colletotrichum gloeosporioides (O. praelonga) EF593338

99 C. acutatum DQ002836

C. acutatum DQ002847

C. graminicola DQ002859 99 C. graminicola DQ002853

100 C. graminicola DQ002860 100 C. sublineolum DQ002869

74 C. coccodes DQ002829

C. magna DQ002828

C. fragariae DQ002827

C. gloeosporioides DQ002825

C. musae DQ002817 76 G. cingulata AY357890

C. gloeosporioides DQ002816

C. gloeosporioides DQ002823

FIG. 3.6. Phylogenetic tree (Inferred MP tree) among Colletotrichum spp. and Colletotrichum sp. strains isolated from Fioriia externa, for the High Mobility Box at the MAT 1-2 gene. Consensus of two equally parsimonious trees. Numbers above nodes are bootstrap values based of 1000 iterations with 30 random additions each. Only branches with values above 70 are considered well supported.

110 TABLE 3.1 – Characterization of different Colletotrichum spp.

Rank Fungi Identification Geographic origin Year Host Code score (*) 28S ITS B-tubulin2 GPDH GS MAT1-2 th Entomopathogen Colletotrichum sp. Mohonk, NY 2005 F. externa EHS36 5 EF464578 EF464591 EF593320 EF593339 EF593348 EF593357 rd Colletotrichum sp. Mohonk, NY 2005 F. externa EHS41 3 EF464579 EF464592 EF593321 EF593340 EF593349 EF593358 th Colletotrichum sp. Bayberry Lane, NY 2005 F. externa EHS48 13 EF464580 EF464593 EF593322 EF593341 EF593350 EF593359

rd Colletotrichum sp. Esopus, NY 2005 F. externa EHS51 23 EF593331 EF593369 EF593323 EF593342 EF593351 EF593360

th Colletotrichum sp. Esopus, NY 2005 F. externa EHS52 8 EF593332 EF593370 EF593324 EF593343 EF593352 EF593361 st Colletotrichum sp. Ward Pound Ridge 2005 F. externa EHS58 1 EF464581 EF464594 EF593325 EF593344 EF593353 EF593362 Res., NY

rd Colletotrichum sp. Ward Pound Ridge 2005 F. externa EHS61 23 EF464582 EF464595 EF593326 EF593345 EF593354 EF593363 Res., NY 111 ∆ C. gloeosporiodes Rio de Janeiro, BZ 1994 O. praelonga ARSEF4360 - EF593337 EF593371 EF593327 EF593346 EF593355 EF593366 C. gloeosporiodes Sao Paulo, BZ 2006 O. praelonga ∇ - EF593338 EF593372 EF593328 EF593347 EF593356 EF593367 EMA26

TM EF593333 EF593329 EF593368 Phytopathogen C. acutatum TopCrop , NJ 2006 Blueberry fruit ERL1379 - (Vaccinium sp.)

C. acutatum Burlington, VT 2006 Tomato fruit ERL1380 - EF593334 EF593330 (Solanum lycopersicum)

Colletotrichum sp. Bayberry Lane, NY 2006 Tulip tree leaf ERL1385 - EF593335 EF593364

Colletotrichum sp. Bayberry Lane, NY 2006 Apple fruit var. golden ERL1395 - EF593336 EF593365 delicious

(*) Non-parametric ranking comprising overall strain performance based on germination rate, speed of growth and conidia production (Parker et al 2005). (∆) Obtained from the USDA Entomopathogenic Fungal Collection, Ithaca, NY. (∇) Obtained from EMBRAPA Colletotrichum gloeosporioides f. sp. ortheziidae, Jaguariúna, São Paulo, Brazil. () Sequencing not performed. () No amplification obtained with primers HMGacuF and HMGacuR.

111 TABLE 3.2. Morphological plasticity in Colletotrichum sp. isolated in epizootic areas

Inoculum source – SUSPENSION

Character state

Fungal pigmentation◊ Conidia◊ Mycelium◊ Media pigmentation◊ morphology

Temp. G O W P R B C C+ S A B O G P Cr. W

10 C 84 21.3 ------98.6 1.3 1.4 17.3 81.3 - - -

15 C 86.6 32 1.3 - - - - 8 89.4 10.6 4 25.4 76 16 2.6 -

20 C 85.3 37.3 - 32 - - - 16 97.3 2.7 16 49.4 42.6 53.4 34.7 -

25 C 29.4 18.6 1.3 80 4 6.7 9.4 89.4 10.6 - 20 13.3 42.6 42.6 -

Inoculum source – SINGLE SPORE

Temp. G O W P R B C C+ S A B O G P Cr. W

10 C 90.6 10.6 - - - 13.3 - - 94.6 5.4 89.3 62.6 10.6 - - 92

15 C 100 5.3 5.3 - - 2.6 - - 90.7 9.3 16 40 1.3 53.3 - 41.3

20 C 92 5.3 - 72 - - - - 89.4 10.6 15.5 5.3 5.3 22.6 61.3 5.3

25 C 92 33.4 8 65.3 - - - - 98.7 1.3 2.6 4 2.6 - 97.3 -

◊ All values are percentage of total number of plates presenting the character state (N=78, 3 replicates were tested for each temperature); Conidia masses: Scattered (C); Uniformly distributed (C+); Mycelium morphology: Aerial (A); Superficial (S); Mycelium and medium pigmentation: Black (B); Gray (G); Aerial dense gray (G+); Orange (O); Pink (P); Red (R); White (W); Chromogenic (Cr.).

112 TABLE 3.3. Cross-fertile bioassay in toothpick substratum

Toothpick crosses

Region Isolate EHS36 EHS37 EHS41 EHS47 EHS48 ERL 1385 ERL 1395 EHS50 EHS51 EHS52 EHS56 EHS58 EHS61 ARSEF4360

EHS36 Nt - + +/- - - + + + + + + + Mohonk, NY EHS37 - Nt - + + - - - - - +/- - + EHS41 + - Nt - + +/- Nt +/- +/- + +/- +/- +/- +

EHS47 + +/- - Nt + - Nt +/- - +/- - - - - Bayberry Lane, EHS48 +/- + + + Nt +/- - + + + + + + - NY Tulip ERL 1385 - - +/- - +/- Nt +/- +/- - + + +/- +/- +/- Apple ERL 1395 - - Nt Nt - +/- Nt Nt - - - - Nt +/-

EHS50 + - +/- +/- + +/- Nt Nt + - + - +/- +/- Esopus, NY EHS51 + - +/- - + - - + Nt - + - - + EHS52 + +/- + +/- + + - - - Nt - - - -

113

Ward Pound EHS56 + - +/- - + + - + + - Nt - - - Ridge, NY EHS58 + +/- +/- + + +/------Nt +/- + EHS61 + - +/- - + +/- Nt +/- - - - +/- Nt +/-

Jaguariuna, Sao Brazil + - + - - +/- +/- +/- + - - + +/- Nt Paulo, BR ARSEF4360

+ /- = few stalled structures; + = many stalled structures; - = No structures; Nt = Not tested.

113 TABLE 3.4. Conidia morphology of entomopathogenic Colletotrichum spp. at 22 C in PDA medium examined in water and stained with cotton blue

Conidial length × width (µm)

Age Shape Mean Range Area Strain□ N SD (d) factor (µm) 95% Confidence Interval (µm2)

Colletotrichum sp. from US epizootic sites

A,B D-F D-G D,E EHS35 14 51 0.67 6.63 × 3.14 [ 6.15-7.11 × 2.94-3.35 ] 1.71 × 0.72 21.05 A E,F F-H D,E EHS36 14 50 0.73 5.61 × 2.91 [ 4.78-6.43 × 2.55-3.28 ] 2.90 × 1.28 18.61 A,B D-F C-E C-E EHS37 14 52 0.70 6.60 × 3.44 [ 6.06-7.13 × 3.24-3.64 ] 1.91 × 0.71 23.10 A,B D-F D,C C-E EHS39 14 55 0.71 6.85 × 2.43 [ 6.19-7.51 × 3.36-3.75 ] 2.43 × 0.72 24.59 A,B D-F E-H D,E EHS40 14 51 0.66 6.63 × 2.99 [ 6.25-7.01 × 2.84-3.15 ] 1.35 × 0.54 19.71 A,B C A,B A EHS41 14 52 0.67 8.57 × 3.95 [ 7.22-9.91 × 3.27-4.63 ] 4.82 × 2.44 43.78 A,B E,F D,C C-E EHS42 14 50 0.72 6.18 × 3.52 [ 5.63-6.73 × 3.29-3.75 ] 1.94 × 0.81 22.30 A,B E,F C-F D,E EHS43 14 50 0.71 6.15 × 3.38 [ 5.69-6.61 × 3.15-3.62 ] 1.62 × 0.82 20.84 114 A,B D-F B,C C-E EHS44 14 56 0.70 6.94 × 3.72 [ 6.38-7.49 × 3.50-3.93 ] 2.05 × 0.80 25.59 A,B D-F D-H D,E EHS45 14 99 0.68 6.56 × 3.08 [ 6.09-7.02 × 2.93-3.23 ] 2.32 × 0.74 20.68 A,B D,E A B,C EHS46 14 81 0.73 7.15 × 4.13 [ 6.69-7.62 × 3.98-4.28 ] 2.09 × 0.68 29.75 A,B D-F C-E C-E EHS47 14 58 0.70 6.65 × 3.49 [ 6.26-7.04 × 3.32-3.66 ] 1.48 × 0.65 23.13 A,B D-F C-F C-E EHS48 14 65 0.66 7.10 × 3.27 [ 6.58-7.61 × 3.08-3.47 ] 2.06 × 0.77 22.65 A,B D-F C-F D,E EHS49 14 51 0.69 6.54 × 3.33 [ 6.00-7.08 × 3.16-3.51 ] 1.93 × 0.62 21.39 A,B E,F D-H D,E EHS50 14 57 0.71 6.13 × 3.06 [ 5.64-6.61 × 2.91-3.21] 1.82 × 0.57 19.29 A,B E,F G-I E EHS51 14 54 0.67 6.27 × 2.80 [ 5.69-6.85 × 2.63-2.98 ] 2.12 × 0.64 17.34 A,B E,F G-I E EHS52 14 99 0.66 6.26 × 2.78 [ 5.94-6.57 × 2.69-2.87 ] 1.68 × 0.47 17.26 A,B D,C A B EHS53 14 52 0.72 7.85 × 4.22 [ 7.02-8.68 × 3.99-4.45 ] 2.97 × 0.81 33.45 A,B D-F C-F C-E EHS55 14 60 0.67 7.06 × 3.32 [ 6.70-7.41 × 3.18-3.47 ] 1.36 × 0.55 23.31 B E,F H-I E EHS56 14 55 0.65 6.38 × 2.73 [ 5.89-6.87 × 2.58-2.88 ] 1.82 × 0.54 17.31 A,B D-F C-F D,E EHS57 14 64 0.71 6.44 × 3.28 [ 5.97-6.91 × 3.15-3.42 ] 1.88 × 0.55 21.64 A,B D- C-G D,E EHS58 14 58 0.69 6.64 × 3.24 [ 6.00-7.28 × 3.05-3.43 ] 2.44 × 0.72 21.40 A,B D-F D,C C-E EHS59 14 51 0.69 6.75 × 3.57 [ 6.08-7.43 × 3.31-3.84 ] 2.40 × 0.94 22.97 A,B E,F C-F D,E EHS61 14 79 0.71 6.01 × 3.29 [ 5.64-6.37 × 3.13-3.45 ] 1.61 × 0.71 19.56 A,B E,F D-H D,E EHS63 14 75 0.71 5.96 × 3.14 [ 5.55-6.36 × 2.99-3.29 ] 1.76 × 0.63 18.55 A,B D-F D-H D,E EHS66 14 54 0.67 6.80 × 3.10 [ 6.18-7.41 × 2.88-3.31 ] 2.24 × 0.79 20.45

(Continue)

114 TABLE 3.4 (Continued). Conidia morphology of entomopathogenic Colletotrichum spp. at 22 C in PDA medium examined in water and stained with cotton blue

Conidial length × width (µm)

Age Shape Mean Range Area Strain□ N SD (d) factor (µm) 95% Confidence Interval (µm2)

■ Range 0.65-0.73 5.61-8.57 × 2.73-4.22 [ 4.78-9.91× 2.55-4.63 ] 1.35-4.82 × 0.47-2.44 17.26-43.78 Mean 0.69 6.62 × 3.31 [ 6.51-6.73 × 3.26 -3.35] 2.22 × 0.91 22.42

Colletotrichum gloeosporioides from Brazilian epizootic sites

C B I,J C-E EMA26 14 99 0.49 10.35 × 2.50 [ 9.80-10.92 × 2.39-2.62 ] 2.76 × 0.55 26.17 D A J C,D ARSEF4360 14 73 0.44 11.88 × 2.37 [11.11-12.66 × 2.23-2.52 ] 3.33 × 0.62 27.31

115

□ Values with different letters are statistically different according to the Student-Newman Keuls (SNK) post-hoc test performed using SAS®

■ Reported range of means (length x width) for C. acutatum in PDA medium at 21-25 C ; examined in water = 13.2-15.9 × 4.4-4.9 µm.

C. gloeosporioides (self-sterile strains) = 15.0 -16.6 ×5.3-5.5 µm; C. gloeosporioides (self-fertile strains) = 14.8 -16.3 × 5.4-5.5 µm.

(Guerber and Correll 2001).

■ Reported range of means (length x width) for C. acutatum in PDA medium at 22 C, examined in water = 9.3-16.9 × 3.1-5.4 µm.

C. gloeosporioides = 14.7-18.8 × 5.0-6.2 µm.

(Du et al 2005).

115 TABLE 3.5. Appressoria morphology of entomopathogenic Colletotrichum sp. strains growing in strawberry leaves at 22 C examined in water and stained with cotton blue

Appressoria length × width (µm)◊

Shape Mean Range Area Strain□ N Perimeter SD factor (µm) 95% Confidence Interval (µm2)

Colletotrichum sp. from US epizootic sites

A-E A A-C A-C A EHS35 53 0.80 23.20 7.80 × 6.75 [ 7.47-8.14 × 6.49-7.01 ] 1.19 × 0.94 34.31 A,B D,E E-G D-H F-H EHS36 48 0.83 19.67 6.82 × 5.91 [ 6.63-7.02 × 5.66-6.16 ] 0.67 × 0.85 25.68 B-F E J H J EHS37 48 0.77 19.87 6.35 × 5.58 [ 6.18-6.52 × 5.40-5.76 ] 0.57 × 0.65 22.46 B-F C-E D-I C-H F-H EHS39 49 0.78 20.56 7.03 × 6.11 [ 6.80-7.26 × 5.79-6.43 ] 0.81 × 1.11 25.66 A-D C-E D-I B-H D-H EHS40 38 0.80 20. 52 7.13 × 6.16 [ 6.88-7.37 × 5.88-6.44 ] 0.74 × 0.85 27.84 A-D E G-J G,H I,J EHS41 63 0.81 19.05 6.73 × 5.62 [ 6.54-6.92 × 5.41-5.84 ] 0.75 × 0.85 23.49 B-F D,E H-J E-H H-J 116 EHS42 46 0.79 19.58 6.68 × 5.87 [ 6.47-6.89 × 5.64-6.09 ] 0.69 × 0.75 24.22 A-D D,E G-F D-H G-J EHS43 58 0.81 19.59 6.76 × 5.91 [ 6.55-6.97 × 5.69-6.13 ] 0.79 × 0.84 24.78 B-F C-E B-H A-G F-H EHS44 54 0.77 20.70 7.31 × 6.29 [ 6.99-7.63 × 5.90-6.67 ] 1.16 × 1.40 26.20 B-F D,E I-J F-H H-J EHS45 55 0.78 19.68 6.61 × 5.69 [ 6.40-6.81 × 5.51-5.86 ] 0.74 × 0.64 24.22 A-C A,B A,B A,B A,B EHS46 35 0.82 22.52 7.85 × 6.85 [ 7.52-8.17 × 6.47-7.23 ] 0.95 × 1.10 33.32 A B-D D-I B-H B-D EHS47 61 0.85 21.22 7.06 × 6.20 [ 6.92-7.21 × 6.86-6.33 ] 0.56 × 0.51 30.50 F B-D D-I B-H F-H EHS48 59 0.74 21.23 7.14 × 6.21 [ 6.80-7.47 × 5.89-6.54 ] 1.29 × 1.25 26.38 B-F B-D B-H B-H D-G EHS49 42 0.79 21.03 7.29 × 6.18 [ 7.02-7.56 × 5.90-6.46 ] 0.85 × 0.90 28.06 B-F A,B A-D A-E B,C EHS50 66 0.77 22. 59 7.61 × 6.53 [ 7.36-7.86 × 6.29-6.77 ] 1.01 × 0.98 31.51 A-D C-E C-I D-H D-H EHS51 78 0.81 20.58 7.20 × 6.03 [ 6.99-7.42 × 5.81-6.25 ] 0.94 × 0.96 27.44 B-F A-C B-D A-E C-E EHS52 49 0.79 21.76 7.53 × 6.50 [ 7.28-7.77 × 6.19-6.81 ] 0.85 × 1.07 30.08 A,B D,E D-I D-H F-H EHS53 53 0.83 19.79 7.06 × 5.96 [ 6.87-7.25 × 5.76-6.16 ] 0.68 × 0.73 26.06 B-F B-D C-I A-F D-H EHS55 48 0.78 21.07 7.17 × 6.34 [ 6.88-7.45 × 6.11-6.57 ] 0.97 × 0.80 27.77 A-D B-D B-F A-G C-F EHS56 53 0.81 21.10 7.42 × 6.33 [ 7.20-7.63 × 6.05-6.60 ] 0.78 × 0.99 28.78 C-F A-C B-G A-H C-F EHS57 62 0.77 21.75 7.37 × 6.26 [ 7.18-7.55 × 6.04-6.48 ] 0.72 × 0.87 28.83 B-F B-D C-I A-E D-H EHS58 50 0.77 21.29 7.18 × 6.54 [ 6.90-7.46 × 6.22-6.86 ] 0.99 × 1.12 27.78 E-F B-D C-I D-H E-I EHS59 52 0.75 21.26 7.19 × 6.02 [ 6.87-7.51 × 5.66-6.39 ] 1.14 × 1.30 26.82 D-F B-D B-D A-F D-G EHS61 11 0.76 21. 36 7.47 × 6.39 [ 6.84-8.09 × 5.53-7.25 ] 0.92 × 1.27 28.07

(Continue)

116 TABLE 3.5 (Continued). Appressoria morphology of entomopathogenic Colletotrichum sp. strains growing in strawberry leaves at 22 C

examined in water and stained with cotton blue

Appressoria length × width (µm)◊

Shape Mean Range Area Strain□ N Perimeter SD factor (µm) 95% Confidence Interval (µm2)

B-F A-C B-D A-D B,C EHS63 72 0.79 22.22 7.50 × 6.62 [ 7.31-7.69 × 6.42-6.82 ] 0.80 × 0.85 31.45 B-F A-C A-D A-G C-E EHS66 43 0.79 21.77 7.58 × 6.31 [ 7.34-7.83 × 6.03-6.59 ] 0.79 × 0.90 29.95

■ Range 0.74-0.85 19.05-23.20 6.35-7.85 × 5.58-6.85 [ 6.18-7.47 × 5.40-5.76 ] 0.57-1.29 × 0.51-1.40 22.46-34.31 Mean 0.79 20.96 7.1 × 6.19 [ 6.35-7.85 × 5.58-6.85 ] 0.94 × 0.99 27.73

117 Colletotrichium gloeosporioides from Brazilian epizootic sites

G A A A B-D EMA26 41 0.71 23.03 8.11 × 6.90 [ 7.60-8.62 × 6.46-7.35 ] 1.61 × 1.41 30.38 ARSEF4360 ------

◊ Measurements were made after 24 h.

□ Values with different letters are statistically different according to the Student-Newman Keuls (SNK) post-hoc test performed using SAS®

■ Reported range for C. acutatum appressoria (length x width) produced in deionized H2O, in a moist chamber overnight = 5.8-10.3 × 4.6-9.5 µm;

C. gloeosporioides = 5.7-8.8 × 4.2-6.6 µm.

(Du et al. 2005).

117 TABLE 2.6. Perithecia of Glomerella sp. on snap bean stems (2 cm) at 22 C from cross-fertile ERL1385 x EHS58

Tri-dimensional structure Perithecia length × width (µm) Form Samples□ Elliptical Shape factor Mean Area N (%) SD Factor (µm) (µm2) ◦ ◦ ◦ A G O

◊ A A A A A A A A Rep. 2 26 1.20 0.83 33 46 21 115.39 × 108.83 29.62 ×29.75 8639.28

♦ A B B A B B Rep. 2-a 14 1.09 0.62 64 36 0 198.68 B × 183.75 B 98.36 × 92.06 32211.60

■ Mean 1.16 0.75 146.08 × 136.43 74.85 × 69.78 17323.82

118 Measurements for self-fertile sample were not estimated due to copious saprophytic growth in the sample.

◊ Perithecia produced in bean stem retrieved from the plant after three wk inoculation and seven d on MSM;

♦ Perithecia retrieved directly from the plant stem;

□ Values with different letters are statistically different according to the Student-Newman Keuls (SNK) post-hoc test performed using SAS® ;

■ Reported range of mean width for G. acutata retrieved from crosses in birch toothpicks on minimal salts medium (MSM), at 22 C, and after

26-41 d, under constant fluorescent light, and examined in water = 156-203 µm (Guerber and Correll 2001); ◦ Perithecium form: A= Ampulliform; G= Globose; O=Ovoid.

118 TABLE 3.7. Ascospores of Glomerella sp. on snap bean stems (2 cm) at 22 C and under natural light

Ascospore length × width (µm)

Samples□ N Shape factor Mean Range SD Area (µm) 95% Confidence Interval (µm2)

A A Rep.2 64 0.47 9.12 A × 3.16 A [ 8.78-9.45 × 3.05-3.28 ] 1.34 × 0.46 39.29 B B Rep.2-a 55 0.57 9.17 A × 3.02 A [ 8.84-9.50 × 2.91-3.13 ] 1.22 × 0.40 28.30

■ Range 0.47-0.57 9.12-9.17 × 3.02-3.16 [ 8.78-9.50 × 2.91-3.28 ] 1.34-1.22 × 0.40-0.46 28.30-39.29

Mean 0.51 9.14 × 3.10 [ 8.91-9.38 × 3.02-3.18 ] 1.28 × 0.44 34.21

119

Measurements for self-fertile sample were not estimated due to copious saprophytic growth in the sample.

□ Values with different letters are statistically different according to the Student-Newman Keuls (SNK) post-hoc test performed using SAS®;

■ Reported range of means (length x width) for G. acutata retrieved from crosses in birch toothpicks; on MSM; after 26-33 d; at 20 C

under constant fluorescent light and examined in water = 12.2-17.7 × 5.0-6.5 µm (Guerber and Correll 2001).

119 CHAPTER 4

Host Plant Associations with an Entomopathogenic Strain of Colletotrichum acutatum from Elongate Hemlock Scale

José A. P. Marcelino1a, Svetlana Gouli1, Bruce L. Parker1, Margaret Skinner1 Rosanna Giordano2 and Lora Schwarzberg3

1 Entomology Research Laboratory, University of Vermont (UVM), Burlington, Vermont, 05405-0105 USA 2 Illinois Natural History Survey, Division of Biodiversity and Ecological Entomology, Champaign, Illinois, 61820 USA 3 New York State Department of Environmental Conservation, Albany, New York, 12233-1750 USA

Abstract

A fungal epizootic has been detected in populations of Fiorinia externa in hemlock forests of several northeastern states. Colletotrichum acutatum, a well-known plant pathogen, was the most commonly recovered fungus from infected scales. This is the second report of a Colletotrichum sp. infecting scale insects. In Brazil C. gloeosporioides f. sp. ortheziidae recovered from Orthezia praelonga is under development as a biopesticide for citrus production. The potential of developing the Fiorinia strain for management of F. externa is being evaluated. This fungus was also detected growing endophytically in 28 species of plants within the epizootic areas. Using molecular methods, it was determined that the Colletotrichum strains recovered from scales (C. acutatum f. sp. fiorinia) and plants were identical for the High Mobility Box at the MAT1-2, mating type gene. Results from plant bioassays confirmed that this entomopathogenic Fiorinia strain exhibits the capacity to grow endophytically in several of the plants tested without causing external symptoms or signs of infection. In addition, this strain causes mild symptoms of infection in strawberry plants. The implications of these findings for the use of this fungus as a biological control agent are discussed.

Keywords: Fiorinia externa, Colletotrichum sp., epizootic, elongate hemlock scale, host range Corresponding author: a [email protected]

120 Introduction

The eastern hemlock, Tsuga canadensis (L.) Carrière, a common species in forests in the northeastern United States, is showing evidence of significant decline in some areas. One causal agent associated with this decline is Fiorinia externa Ferris

(Hemiptera: Diaspididae), elongate hemlock scale, an exotic invasive from Japan accidentally introduced in the early 1900s (Sasscer 1912, Lambdin et al. 2005). Chemical insecticides and classical biological control methods have proven impractical and provided inconsistent efficacy (Smith and Lewis 2005). Considering the value of eastern hemlock to forest biodiversity, effective methods to manage F. externa are needed.

In 2002 an epizootic was reported within a population of F. externa in the Mianus

River Gorge Preserve in Bedford, NY and attributed to unidentified fungi which produced sclerotia that often completely covered the insect (McClure 2002). Several additional epizootic sites have been identified in 36 different geographical localities in

New York, Pennsylvania, Connecticut and New Jersey. The most prevalent fungus isolated from these epizootics has been morphologically and molecularly identified as

Colletotrichum acutatum f. sp. fiorinia, the Fiorinia strain (unpublished data) and has shown the ability to be easily cultured in vitro (Parker et al. 2005).

Though C. acutatum is more commonly known as a phytopathogen (Bailey and

Jeger 1992, Prusky et al. 2000), one report was found of C. gloeosporioides f. sp. ortheziidae causing epizootics in the scale Orthezia praelonga Douglas 1891 (Hemiptera:

Ortheziidae), a major pest of Citrus spp. in Brazil. Research on the biological control of this pest using the Orthezia strain has been conducted in Brazil for >13 yr and this strain

121 is under commercial development (Cesnik and Ferraz 2000, R. Cesnik, pers. comm.

2006). Infection of F. externa by the Fiorinia strain represents the second reported case

of a member of this genus infecting a scale insect.

The distribution and infective capacity of the Fiorinia strain must be assessed to

evaluate the biological control potential of this organism. An assessment of its natural

occurrence in plants in the hemlock forest ecosystem and its phytopathogenicity to

several horticultural crops were made. This information is critical before further

development of the Fiorinia strain for use against F. externa can be considered.

Material and Methods

Plant bioassays

Isolates. Several entomopathogenic and phytopathogenic fungi were tested to

determine their ability to infect horticultural plants and eastern hemlock seedlings (Table

4.1). The following strains of C. acutatum were tested: five entomopathogenic strains of

C. acutatum f. sp. fiorinia (hereafter called Fiorinia strains) obtained from pure culture

lines isolated from F. externa in sites where the epizootic occurs; and two

phytopathogenic C. acutatum strains, one from a blueberry fruit (ERL1379) and one from a

tomato fruit (ERL1380). To compare the phytopathogenicity of the Fiorinia strains with a recognized entomopathogen, one strain of Lecanicillium lecanii (Zimmerman) Gams &

Zare [= Verticillium lecanii (Zimm.) Viégas] (EHS132) isolated from F. externa was

included in the bioassays. In the 2006 bioassays, the entomopathogenic C.

gloeosporiodes. f. sp. ortheziidae (ARSEF4360) from Brazil was also included. Isolates

were grown in potato dextrose agar (PDA) (39 g/l) supplemented with penicillin (5 ml/l)

122 and streptomycin (12.5 ml/l) for 10-12 d before being harvested with sterile Pasteur pipettes to obtain suspensions of the isolates, in sterile distilled water (SDW), for

subsequent calibration of conidial concentrations.

Table 4.1. Fungal isolates used for the horticultural plant and hemlock phenological bioassays

Fungus Year of Species Code Host Geographic origin type collection

Entomo- Colletotrichum f. sp. EHS41 Fiorinia Mohonk, NY 2005

pathogenic fiorinia (Fiorinia externa

fungi strain)

Fiorinia strain EHS48 F. externa Bayberry Lane, NY 2005

Fiorinia strain EHS51 F. externa Esopus, NY 2005

Fiorinia strain EHS58 F. externa Ward Pound Ridge 2005

Reservation, NY

Fiorinia strain EHS61 F. externa Ward Pound Ridge 2005

Reservation, NY

Lecanicilium lecanii EHS132 F. externa South Salem, NY 2005

C. gloeosporiodes ARSEF4360 Orthezia Jaguariuna, Sao 1994

f. sp. ortheziidae praelonga Paulo, Brazil

Phyto- C. acutatum ERL1379 Blueberry NJ 2005

pathogenic fruit

fungi C. acutatum ERL1380 Tomato fruit Burlington, VT 2005

Plants. The virulence of the above fungal isolates was tested on the following horticultural crops: pepper (Capsicum annuum var. New Ace, Solanaceae), tomato

123 (Solanum lycopersicum var. Patio, Solanaceae); common bush snap bean (Phaseolus

vulgaris var. Blue Lake 274, Fabaceae); strawberry (Fragaria x ananassa var. Honeoye,

Rosaceae) and barley (Hordeum vulgare, Gramineae). These plant species were selected

for testing because they are reported to be highly susceptible to C. acutatum (Ivey et al.

2004, Dillard and Cobb 1998, Tu 1992, Horowitz et al. 2004, Martin and Skoropad

1978). All plants were grown in a greenhouse from seed in 23-cm diam pots containing

Metro-Mix 360 potting medium (Sungro Horticulture, Vancouver, Canada). Four wk old

plants were used for these bioassays. In addition to the horticultural plants, 4 yr old

eastern hemlock seedlings (Western Maine Nurseries, Fryeberg, ME) were also tested.

Only plants with no external signs of fungal infection or disease were selected for testing.

Each plant was sprayed individually to runoff with 30 ml of a suspension of 106 conidia/ml-1 SDW containing 0.02% Silwet L-77 as a wetting agent (Loveland Industries,

Inc., Greeley, CO). Suspensions were calibrated to the correct conidial concentration per

ml SDW with an Improved Neubauer haemacytometer (Propper®) according to the

protocol of Goettel and Inglis (1997). Suspensions were applied with a hand-held air-

brush sprayer (Badger Air-Brush Co., Franklin Park, IL) operating at 1.75 kg/cm2 in a

sterile fume hood. Controls were treated with SDW containing 0.02% Silwet L-77. After

treatment, plants were incubated in a dew chamber (3.95 m high × 1.10 m wide × 1.10 m

deep built with 2.5 cm diam PVC pipe and covered with plastic) for 24 h at 22 ± 1oC

according to protocols of TeBeest (1988), then transferred to a naturally illuminated

greenhouse with adjusted temperature (22 ± 0.5oC), and arranged on benches in a

124 completely randomized block design. Each treatment was replicated four times per plant species, and the complete bioassay was run once in 2005 and 2006.

Koch’s postulates were performed 1 mo after treatment. Four to six leaf samples

(approx. 2 sq. cm) were excised from each plant and surface sterilized by immersion in

75% ethanol + 0.02% Silwet L-77 for 20 s (Cowles et al. 2000), followed by 5 s in SDW,

45 s in 2.5% sodium hypochlorite (NaOCl) and finally rinsed twice in SDW for 10 s.

Samples were air dried and placed on PDA supplemented with penicillin (5 ml/l) and

streptomycin (12.5 ml/l). Cultures were incubated in the dark at 22oC for 10 d after which

re-isolation of the test fungi was attempted.

Hemlock phenological trials. The potential impact of the test fungi on eastern

hemlock during the growing season was assessed using 1.5-yr old seedlings (Intervale

Conservation Nursery, Burlington, VT) and 4.5-yr old seedlings (Western Maine

Nurseries, Fryeberg, ME). Seedlings were transplanted into Metro-Mix 360 in 11.5-, and

23-cm diam pots, for the 1.5 and 4.5-yr old seedlings, respectively, and grown outside

prior to treatment.

To assess changes in potential infectivity of the fungi over the growing season, a

new group of 40 seedlings (4 plants per treatment and seedling age) were treated each

month from June to September for the 1.5-yr old seedlings and May to September for the

4.5-yr old seedlings. Fungal treatments were applied as described above. After treatment,

seedlings were incubated in a dew chamber for 24 h at 22 ± 0.5oC and then held in a

greenhouse with ambient light and temperature ranging from 9.8 to 24oC depending on

the month. Temperature was monitored using HOBO Data Loggers (Onset Computer

125 Corporation, Bourne, MA). Following treatment, seedlings were inspected monthly for symptoms or signs of disease. A single bioassay was performed during the 2006 plant growing season.

Koch’s postulates were performed 1 mo after treatment as described above, sampling eight needles from a randomly selected twig per seedling. This was repeated at monthly intervals throughout the test period.

Statistical analyses. For the horticultural plant assays, the susceptibility to fungal infection for all test plants was rated according to the probability of re-isolating all fungal strains following treatment. The strawberry plants were used as a reference for infection because this plant is highly susceptible to infection by C. acutatum (Horowitz et al.

2004). The frequency of recovering a given fungal strain was also determined for all plant types. The phytopathogenic ERL1380 were selected at random as a reference

Colletotrichum sp. isolate to which other test isolates were compared.

All data from the bioassays were analyzed using a logistic regression analysis for binary variables. Plant species tested in the bioassays were treated as covariates. Adjusted

(log10) odds ratios (with 95% confidence intervals) were calculated using SAS (SAS

Institute 1990) and plotted using GraphPad software (Motulsky 1999). A Wald Chi- square test was used to determine significant differences between variables (SAS Institute

1990). Treatments for which the Fiorinia strain was not re-isolated were excluded from the odds ratio analysis as zeros in the denominator would result in an undefined number.

126

Understory plant screening and molecular identification

Samples of several common plant species were taken in 10 sites where the F.

externa epizootic occurred to determine the presence of C. acutatum growing in or infecting plants. A total of 97 plants representing 50 species growing in different strata in the hemlock forest were sampled, including low-growing shrubs, vines and trees (Table

4.2). Both live-and dead plant material from the litter were sampled. A 15-20 cm branch or stem sample was taken from each live plant. For the leafy plants, two to three leaves were selected at random, from which four pieces (2 cm2) were excised and placed in

individual Petri dishes. For the hemlock needles, eight individual needles (from the litter

or live trees) were sampled and placed in separate dishes. All samples were held in an incubator at (22 ± 0.5oC) prior to processing. Samples were surface sterilized as described above and placed on PDA supplemented with penicillin (5 ml/l) and streptomycin (12.5 ml/l), held in the dark at (22 ± 0.5oC) for 7 d and then inspected for the presence of the Fiorinia strain. This strain was distinguished from other fungi based on the following characteristics: mycelium and growth medium were pinkish or intensely red, conidia were the typical shape and size for that strain, and the fungus grew faster than other fungi.

To confirm that the C. acutatum from the plant material was the entomopathogenic Fiorinia strain, a sub-sample of 20 strains recovered from 19 different

plant species were identified by molecular methods. DNA was extracted from 1-wk old cultures using the Power SoilTM DNA kit (Mo Bio Laboratories, Inc., Carlsbad, CA)

following the manufacturers’ directions with the following exceptions: 1) Samples were

127 shaken for 5 min at 5.5 m/s to facilitate breakage of the cell walls using a FastPrepTM

FP120 machine (Thermo Savant, Holbrook, NY); 2) DNA was eluted using 100 µl of diluted elution buffer (1:15) (Qiagen, Valencia, CA), and concentrated to 20 µl with a speed vacuum (Eppendorf Centrifuge 5415C, Vaudaux, Schönenbuch, Switzerland). The

High Mobility Box (HMG) of the mating-type gene (MAT 1-2) was amplified using primers HMGacuF and HMGacuR (Du et al. 2005).

Polymerase Chain Reaction (PCR) was conducted using Ready-To-Go PCR beads

(Amersham Biosciences Inc., Piscataway, NJ) and the following protocol: initial denaturation at 95 oC for 2 min followed by 30 cycles of 95 oC for 30 s (denaturation), 55

oC for 30 s (annealing) and 72o C for 1 min (elongation). PCR products were purified

using the QIAquick PCR purification kit (Qiagen, Valencia, CA) or Princeton

Separations Centri SpinTM columns (Princeton, Adelphia, NJ). DNA was stored at 4oC.

PCR products were sequenced using a BigDye v3 terminator cycle sequencing kit

(Applied Biosystems, Foster City, CA) and with the following protocol: 95 oC (initial denaturation) for 3 min followed by 30 cycles of 95 oC for 10 s (denaturation), 50 oC for

5 s (annealing), and 60 oC for 2 min (elongation). PCR fragments were sequenced with a

3130xl Genetic Analyzer (Applied Biosystem, Foster City, CA). Chromatograms were

edited and contiguous sequences were generated using SequencherTM (Gene Codes Corp.,

Ann Arbor, MI). Sequences generated from this study were compared with MAT 1-2

gene sequences obtained previously for the Fiorinia strain and available at GenBank.

Primers were removed from sequences. Sequences obtained in this study were also deposited in GenBank® (Table 4.4).

128 Results

Plant bioassays

For all treatments tested, there were no signs of necrosis on plant tissue of pepper,

tomato, beans, barley or hemlock. However, endophytic growth of some of the test fungi

was detected as indicated by re-isolation of the Fiorinia strain from asymptomatic plants

after surface sterilization of leaf samples (Figure 4.1A, Table 4.2). Strawberry leaves

displayed distinct local necrotrophic symptoms and endophytic growth. Though this plant

species appeared to be susceptible to the Colletotrichum sp. treatments, the necrotic spots

(approx. 1 cm diam) that formed did not progress over the entire leaf nor compromise the

viability of the plants 2 mo post treatment (Figure 4.1B).

Figure 4.1. (A) Endophytic growth of Fiorinia strain EHS48 in a bean stem 24 h after treatment, in vitro, showing germinated germ tubes (a) and appressoria (b); (B) Strawberry with necrotic lesions, 2 mo after spraying with the Fiorinia strain.

The percentage of fungal re-isolations per plant species was not significantly

different between test years, allowing data across years to be combined (Wald Chi-square

= 0.49, P = 0.48). Differences in the percentage of plants from which Colletotrichum sp.

129 was re-isolated were significant among the test fungi (Wald Chi-square = 14.72, P =

0.03), and a statistically significant 2-way interaction between the test fungi and plant

species was detected (Wald Chi-square = 71.02, P < 0.0001). Isolate EHS132

(Lecanicilium lecanii) and SDW were excluded from the odds ratio analysis because

Colletotrichum sp. was not re-isolated from these treatments. Approximately 38% of the plants treated with the Fiorinia strains (average of Fiorinia strains reisolation) became infected based on the re-isolation of Colletotrichum sp. (Table 4.2). This was comparable to the phytopathogenic C. acutatum from blueberry (31.3%), but significantly different from the C. acutatum from tomato, and the entomopathogenic Orthezia strain (P<0.05).

Table 4.2. Percentage of fungal re-isolations obtained from the tested plants (N = 8 plants/isolate and plant

species). No fungal re-isolations were obtained from control plants treated with SDW and Silwet

Test plants/ Percent re-isolation Treatment

Overall Barley Beans Hemlock Pepper Strawberry Tomato means

ERL1379 12.5 37.5 12.5 50.0 75.0 0.0 31.3

ERL1380 0.0 75.0 0.0 0.0 75.0 0.0 8.3

ARSEF4360 0.0 0.0 0.0 50.0 0.0 0.0 8.3

EHS132 0.0 0.0 0.0 0.0 0.0 0.0 0.0

EHS41 25.0 50.0 12.5 37.5 62.5 0.0 31.3

EHS48 12.5 50.0 0.0 50.0 75.0 12.5 33.3

EHS51 25.0 87.5 0.0 50.0 50.0 25.0 39.6

EHS58 25.0 75.0 0.0 37.5 87.5 37.5 43.8

EHS61 50.0 75.0 0.0 50.0 75.0 12.5 43.8 SDW 0.0 0.0 0.0 0.0 0.0 0.0 0.0 Overall means 16.7 50.0 2.8 36.1 55.6 9.7

130 The mean odds ratio comparing the number of fungal re-isolations from treated

plants with the reference, ERL1380, for six of the test fungi were >1, indicating that they

were more likely to be re-isolated from the test plants than the reference (Figure 4.2). The

odds ratio for the Orthezia strain was <1, suggesting that it was less likely to be re-

isolated than the other fungi. Because of the variability in the results, differences between

this isolate and four of the others were not significant. The odd ratios demonstrated that

two of the Fiorinia strains were significantly different from the others isolated, indicating

that these two were more infective than the reference isolate.

ERL1379

Fiorinia strain EHS41

Fiorinia strain EHS48

Fiorinia strain EHS51

Fiorinia strain EHS58

Fiorinia strain EHS56 Fungal isolatesFungal tested ARSEF4360

0.01 0.1 1 10

Odds ratio

Figure 4.2. Adjusted odds ratios (95% confidence interval) of endophytic re-isolations obtained with each fungal inoculation treatment in the plant bioassays. The phytopathogenic

Colletotrichum acutatum isolate (ERL1380) from a tomato fruit was used as the reference isolate. An odds ratio of 1 indicated no difference between the probability of re-isolating the test fungus and reference strain; ≥1 indicated a greater probability of re-isolating the test strain than the reference strain; ≤1 indicated a greater probability of re-isolating the reference strain than the test strain. Confidence intervals that overlap the reference line (1) are not significantly different (P = 0.05).

131

The mean odds ratios for test plants were all <1, indicating that it was less likely to recover Colletotrichum sp. from these plants than the reference species (strawberry)

(Figure 4.3). Based on the percentage of plants from which Colletotrichum sp. was re- isolated, indicating endophytic fungal growth, beans and strawberries were the most susceptible, while hemlock and tomato were the least (Table 4.2).

Tomato vs. Strawberry

Pepper vs. Strawberry

Hemlock vs. Strawberry

Beans vs. Strawberry Plants tested Plants

Barley vs. Strawberry

0.001 0.01 0.1 1 10

Odds ratio

Figure 4.3. Adjusted odds ratios (95% confidence interval) of endophytic re-isolations for each plant tested in the plant bioassays. Strawberry was used as the reference plant. An odds ratio of 1 indicated no difference between the probability of re-isolating the fungus from the test plant species and reference plant; ≥1 indicated a greater probability of re-isolating the fungus from the test plant species than the reference species; ≤1 indicated a greater probability of re-isolating the fungus from the reference plant species than the test plants. Confidence intervals that overlap the reference line (1) are not significantly different (P = 0.05).

Hemlock phenological trials

A significant statistical effect across months (Wald Chi-square = 12.73, P = 0.01) and age of plants (Chi-square = 11.53, P < 0.001) for the response of hemlock seedlings

132 to fungi over the growing season was found. The highest percentage of re-isolations

occurred in July among the 1.5 year old seedlings, with between 25-100% recoveries of

Colletotrichum sp. No fungal re-isolations were recovered from 1.5 and 4.5 yr old

hemlock seedlings treated in June, nor in the 4.5 yr old treated in the month of July.

Isolate EHS132 was never re-isolated from hemlock seedlings and excluded from the

analysis (Table 4.3). Based on the odds ratio analysis (Figure 4.4), differences in the rate

of fungal recovery among the test isolates were not significant (Wald Chi-square = 2.92,

P = 0.089).

Table 4.3. Percentage of eastern hemlock seedlings from which Colletotrichum sp. was re-isolated after

treatment (N = 4 seedlings/age and month)

Test month/temperature range (oC)

May1,2 July August September (18.3-22.4) (14.1-21.2) (9.8-18.9) Treatment Seedling age/Percent re-isolation 4.5 yr 1.5 yr 1.5 yr 4.5 yr 1.5 yr 4.5 yr

ERL1379 0 50 50 0 0 0 ERL1380 0 0 50 25 0 0 ARSEF4360 0 0 0 0 0 25 EHS41 0 75 25 0 0 0 EHS48 25 25 0 0 25 0 EHS51 0 50 0 0 0 0 EHS58 0 100 0 0 0 0 EHS61 25 50 0 0 0 0 SDW 0 0 0 0 0 0

1 The 1.5-yr old hemlock seedlings were not available for testing in May.

2 With exception of the month of May, temperatures were monitored with HOBO Data Loggers (Onset Computer Corporation, Bourne, MA).

133

Statistical differences were detected in the susceptibility of the different age of hemlock seedlings tested (Wald Chi-square=11.53, P<0.001). The 1.5-yr old hemlock seedlings were more susceptible to infection than the 4.5-yr old seedlings in the months of July and August. In addition, the timing of application influenced infection, with significantly greater likelihood of infection occurring when plants were sprayed in July than the other months tested. No symptoms of infection or negative impact on seedling growth were observed among the test seedlings.

ERL1379

Fiorinia strain EHS41

Fiorinia strain EHS48

Fiorinia strain EHS51

Fiorinia strain EHS58

Fiorinia strain EHS61 Fungal isolates tested ARSEF4360

0.01 0.1 1 10

Odds ratio

Figure 4.4. Adjusted odds ratios (95% confidence interval) of endophytic re-isolations obtained with fungal treatments in the phenological bioassays. The phytopathogenic Colletotrichum acutatum isolate (ERL1380) from tomato was the reference isolate. An odds ratio of 1 indicated no difference between the probability of re-isolating the reference isolate and the test fungi; ≥1 indicated a greater probability of re-isolating the test strains than the reference isolate; ≤1 indicated a greater probability of re-isolating the reference isolate than the test strains. Confidence intervals that overlap the reference line (1) are not significantly different (P = 0.05).

134

Understory plant screening and molecular identification

Of the 97 plants sampled within areas of the epizootic, 37 plants comprising 28 species in 18 families (52% of sampled species), were found to be positive for the presence of Colletotrichum sp. (Table 4.4). Twenty of these isolates from four locations where the scale epizootic occurs were molecularly identified as the Fiorinia strain (Table

4.4) for the High Mobility Box at the MAT1-2 gene.

Table 4.4. Plant material sampled in epizootic areas and screened for C. acutatum (a sample of live leaves

from one plant per species was taken unless otherwise indicated)

Sample C. acutatum Molecular ID # Plant species (family) sampled Site ◊ recovered GenBank

1 Acer pensylvanicum (Sapindaceae) Negative -

2 A. rubrum (Sapindaceae) Negative -

10 A. pseudoplatanus (Asteraceae) [2 plants] Negative -

2, 9, 10 A. saccharum (Aceraceae) * [3 trees] Endophytic EU043504

2 Alliaria petiolata (Brassicaceae) * Endophytic EU043511

10 Aralia nudicaulis (Araliaceae) Endophytic -

10, 7, 9 Aster sp. (Asteraceae) Endophytic -

1 Arisaema triphyllum (Araceae) * Endophytic EU043499

10 Barberis thunbergii (Berberidaceae) Endophytic -

2 Betula sp. (Betulaceae) Negative -

2 Catalpa speciosa (Bignoniaceae) * Endophytic EU043508

2 Carya sp. (Junglandaceae) Negative -

2 Cornus florida (Benthamidia) Negative -

(Continue)

135 Sample C. acutatum Molecular ID # Plant species (family) sampled Site ◊ recovered GenBank

1 Cypripedium reginae (Orchidaceae) Negative -

1, 4, 8, 10 Dryopteris sp. (Dryopteridaceae) Negative -

2 Fagus grandiflora (Fagaceae) Negative -

1, 3, 5, 8 Fragaria ananassa (Rosaceae) Negative -

10 Hamamelis sp. (Hamamelidaceae) Endophytic -

1 H. virginiana (Hamamelidaceae) * Endophytic EU043501

1 Kalmia latifolia (Ericaceae) Negative -

2 Liriodendrum tulipifera (Magnoliaceae) * [3 trees] Endophytic EU043506

4, 7, 8 L. tulipifera (Magnoliaceae) * [leaves from 5 trees Endophytic -

and litter]

5 L. tulipifera (Magnoliaceae) * [from outside Endophytic EU043516

epizootic]

1 L. tulipifera (Magnoliaceae) * [leaves from 3 trees Endophytic EU043500

and litter]

1 Maianthemum canadense (Ruscaceae) Negative -

6 Malus sp. * (Rosaceae) [fruit and leaves] Signs■ EU043517

2 M. pumila (Rosaceae) * Endophytic EU043509

4 Magnolia sp. (Magnoliaceae) * Endophytic EU043515

2 Myrica rubra (Myricaceae) Negative -

2 Onoclea sensibilis (Onicleaceae) Negative -

10 Pachysandra terminalis (Buxaceae) Endophytic -

2, 10 Parthenocissus quinquefolia (Vitaceae) * Endophytic EU043505

2 Polystichum acrostichoides (Dryopteridaceae) Negative -

2 Prunus cerasifera (Rosaceae) Negative -

(Continue)

136 Sample C. acutatum Molecular ID # Plant species (family) sampled Site ◊ recovered GenBank

2 P. persica (Rosaceae) Negative -

2 P. armenica (Rosaceae) Negative -

3 P. avium (Rosaceae) * Endophytic EU043513

1 Quercus velutina (Fagaceae) Negative -

1 Q. americana (Fagaceae) Negative -

1 Q. alba (Fagaceae) Negative -

2 Q. rubra (Fagaceae) Negative -

2 Rubus sp. (Rosaceae) * Endophytic EU043510

1 R. idaeus (Rosaceae) Endophytic -

10 Rosa sp. (Rosaceae) Endophytic -

2 R. multiflora (Rosaceae) * Endophytic EU043507

1 Sassafras albidum (Lauraceae) * Endophytic EU043498

2 Sorbus americana (Rosaceae) * Endophytic EU043512

2 Symplocarpus foetidus (Araceae) Negative -

10 Taraxacum sp. (Asteraceae) Negative -

1 Thelypteris noveboracensis (Thelypteridaceae) Negative -

2 Tilia americana (Tiliaceae) * Endophytic EU043503

1 Trientalis borealis (Primulaceae) * Endophytic EU043502

9, 10 Tsuga canadensis (Pinaceae) [needles from litter] Endophytic -

9, 10 T. canadensis (Pinaceae) [needles from 4 trees] Negative -

10 Tussilago farfara (Asteraceae) Endophytic -

10 T. farfara (Asteraceae) Negative -

10 Ulmus sp. (Ulmaceae) [2 trees] Endophytic -

3 Vaccinium sp. (Rosaceae) * Endophytic EU043514

(Continue)

137 Sample C. acutatum Molecular ID # Plant species (family) sampled Site ◊ recovered GenBank

10 Verbascum sp. (Scrophulariaceae) Endophytic -

9, 10 Viburnum acerifolium (Adoxaceae) Endophytic -

2, 4, 6, 9,10 V. acerifolium (Adoxaceae) Negative -

* Fungal identification confirmed genetically (High Mobility Box at the MAT1-2, mating type gene).

■ Conidia present in fruit.

◊ Location code (sample date): 1 = Pawling Nature Preserve, Dutchess Co., NY (20 June 2006); 2 = South Salem, Westchester Co., NY (10 Nov. 2005; 5, 16, 20, 27 June 2006); 3 = Lawrence Farm, Orange Co., NY (19 June 2006); 4= Hodgson Farm, Orange Co., NY (19 June 2006); 5 = Whitestone Lane, Rochester City, Monroe Co., NY (13 July 2006); 6 = Poundridge Nurseries, Westchester Co., NY (10 Nov. 2006); 7 = Mountain Lake Park, Westchester Co., NY (10 Nov. 2005); 8 = West Moreland Sanctuary, Westchester Co., NY (11 Jan. 2006); 9 = Olana Historic Site, Columbia Co., NY (29 Aug. 2006); 10 = Macedonia State Park, Litchfield Co., CT (29 Aug. 2006).

These results demonstrate that the Fiorinia strain occurs widely within the epizootic areas, growing endophytically within plants, and remaining viable in dead plant material in the litter.

Discussion

The research reported herein assessed the host plant associations of several isolates recovered within areas where a fungal epizootic occurs in F. externa. The most commonly retrieved fungus from diseased or mummified insects in all sampled sites was molecularly and morphologically identified as C. acutatum f. sp. fiorinia (Fiorinia strain) (Marcelino et al. submitted). This is only the second reported case of a fungus in the genus Colletotrichum, a common phytopathogen, causing an epizootic in an insect

138 pest population. The first was from Brazil where C. gloeosporioides f. sp. ortheziidae

(Ortheziidae strain) is causing epizootics in the scale O. praelonga, a major pest of Citrus spp. (Cesnik and Ferraz 2000). The Fiorinia strain may represent a novel means of sustainable biological control for F. externa in eastern hemlock forests, but an understanding of their infectivity to plants must be determined before they can be considered environmentally safe.

Plant bioassays were conducted to test the infection potential of several Fiorinia strains, the Ortheziidae strain, two C. acutatum strains and one L. lecanii, a known entomopathogen. Several plant species were selected for testing, based on their known susceptibility to Colletotrichum spp. The susceptibility of hemlock seedlings was also tested. With the exception of strawberry, no external signs of fungal infection were observed on the plant species tested. However, evidence of Colletotrichum sp. growing endophytically was commonly obtained, though variation among plant species was observed. Whereas Fiorinia strains were consistently recovered growing endophytically in barley, beans, pepper, strawberry and tomato, only one strain was recovered from hemlock in the plant trials. In contrast, the Ortheziidae strain was recovered only from pepper plants.

Strawberry was found to be the most susceptible to infection by the Fiorinia strains, exhibiting both external necrotrophic and endophytic growth. However, the necrotic lesions occurred sporadically among test plants, and no negative impact on plant growth was observed. In general the necrotic leaf spots did not exceed 1 cm diam and no conidial masses were produced on the leaf surface after 2 mo. Colletotrichum spp.

139 commonly cause anthracnose disease in strawberries and are a serious concern for the strawberry industry (Horowitz et al. 2004). Non-pathogenic strains of this genus have

been reported in strawberries (Horowitz et al. 2002), and it appears that the Fiorinia

strains may be in this category.

In the hemlock phenological bioassays, the Fiorinia strains were commonly

recovered from 1.5 yr old seedlings treated in July. The temperature during this month

(diurnal average 22oC) may have been ideal for infection and colonization of these

strains. These strains and the phytopathogenic C. acutatum strains were recovered

intermittently at other times of the growing season, but no pattern of infection was

observed. Both C. acutatum and C. gloeosporioides have been reported to affect 70 d old

and 4 mo old seedlings of western hemlock, Tsuga heterophylla (Raf.) Sarg. (Lock et al.

1984, Hopkins et al. 1985). However, C. acutatum f. sp. pinea only caused temporary

stunting, and the seedlings recovered within the first year (Canadian Food Inspection

Agency 1997).

Over 90 different plants representing 50 species were sampled in and around the

epizootic areas and processed for infection by Colletotrichum sp. C. acutatum was recovered from 38% of the plants (37 specimens) sampled comprising 28 species in 18 families which represented a broad range of species in the region. Twenty of the isolates recovered were molecularly identified as the Fiorinia strain. These results demonstrate that the Fiorinia strain which has been found to infect F. externa occurs widely within the epizootic areas, growing endophytically within live and dead plant matter.

140 With the exception of strawberry plants, only endophytic growth of the Fiorinia strains was detected in the plants, some of which are common hosts for species in the genus Colletotrichum. We did not observe sporulation either in the field-collected

samples or the plants in the bioassays. It is possible that the Fiorinia strain, which

possesses the capability to infect and kill F. externa, may have additional nutritional

requirements for its growth and reproduction other than plant tissue alone.

Recently it has been suggested that what are considered specialized monomorphic

groups of Colletotrichum may constitute distinct lineages of unknown recent origins that

may inhabit niches other than plants (Zhu and Oudemans 2000, Guerber and Correll

2001, Peres et al. 2005, Crouch et al. 2006). Our results, which report a C. acutatum

infecting F. externa and causing an epizootic, support this hypothesis. It appears that the

C. acutatum f. sp. fiorinia can inhabit niches other than plants, in this case an insect

species. Before the Fiorinia strain can be considered for area-wide use against F. externa

additional research is needed to more fully evaluate its host range, and the conditions

under which it causes infection.

Acknowledgments

We thank Tom Doubleday, Cheryl Frank, Teri Hata and Dr. Robert Jones for their

help with this project. This work was funded in part through a grant awarded by the

Northeastern Area State and Private Forestry, USDA Forest Service (#04-CA-

11244225286) and is in partial fulfillment of requirements for the PhD degree of J.M. at

the University of Vermont.

141 Literature Cited

Bailey JA, Jeger MJ. 1992. Colletotrichum: Biology, Pathology and Control. CAB International. Canadian Food Inspection Agency. 1997. Deregulation of Colletotrichum acutatum, Simmonds. Animal and plant health directorate. Plant protection division. File 3525-11F1-0/3525-11F1/FN7 3510-3-1B1. Available online: http://www.inspection.gc.ca/english/plaveg/protect/dir/d-97-03e.pdf Cesnik R, Ferraz JNG. 2000. Orthezia praelonga, 1891 (Homoptera, Ortheziidae) biologia, controle quimico e biologico. Embrapa Meio Ambiente, Jaguariúna, Brazil, Boletim de Pesquisa 9. 27 p. (in Portuguese) Cowles RS, Cowles EA, McDermott AM, Ramoutar D. 2000. “Inert” formulation ingredients with activity: toxicity of trisiloxane surfactant solutions to twospotted spider mites (Acari: Tetranychidae). J. Econ. Entomol. 93: 180-88. Crouch JA, Clarke BB, Hillman BI. 2006. Unraveling evolutionary relationships among the divergent lineages of Colletotrichum causing anthracnose disease in turfgrass and corn. Phytopathology 96: 46-60. Du M, Schardl CL, Nuckles EM, Vaillancourt LJ. 2005. Using mating-type gene sequences for improved phylogenetic resolution of Colletotrichum species complexes. Mycologia 97: 641-658. Dillard HR, Cobb AC. 1998. Survival of Colletotrichum coccodes in infected tomato tissue and in soil. Plant Disease 82: 235-238. Goettel MS, Inglis GD. 1997. Fungi: Hyphomycetes. In Lacey LA editor. Manual of Techniques in Insect Pathology. Academic Press, London. pp. 213-249. Guerber JC, Correll JC. 2001. Morphological description of Glomerella acutata, the teleomorph of Colletotrichum acutatum. Mycologia 93: 216-229. Hopkins JC, Lock W, Funk A. 1985. Colletotrichum acutatum, a new pathogen on western hemlock seedlings in British Columbia. Canadian Plant Disease Survey 65: 11-13. Horowitz S, Freeman S, Sharon A. 2002. Use of green fluorescent protein-trangenic strains to study pathogenic and non-pathogenic lifestyles in Colletotrichum acutatum. Phytopathology 92: 743-749. Horowitz S, Yarden O, Zveibil A, Freeman S. 2004. Development of a robust screening method for pathogenicity of Colletotrichum spp. on strawberry seedlings enabling forward genetic studies. Plant Disease 88: 845-851. Ivey MLL, Nava-Diaz C, Miller SA. 2004. Identification and management of Colletotrichum acutatum on immature bell peppers. Plant Disease 88 (11): 1198- 1204. Lambdin P, Lynch C, Grant J, Reardon R, Onken B, Rhea R. 2005. Elongate hemlock scale and its natural enemies in the southern Appalachians. In: Onken B, Reardon R editors. 3rd Symposium on Hemlock Woolly Adelgid in the Eastern United States. Asheville, NC 1-3 February 2005 pp. 145-154.

142 Lock W, Sutherland JR, Dennis JJ, Hopkins JC. 1984. Colletotrichum gloeosporioides shoot blight of greenhouse-grown western hemlock seedlings in British Columbia. Plant Disease 68: 628. Motulsky HJ. 1999. Analyzing data with GraphPad Prism. GraphPad Software Inc. San Diego, CA. Martin S, Skoropad W. 1978. Nature of adhesive material of Colletotrichum graminicola appressoria. Transactions British Mycological Society 10: 221-223. McClure MS. 2002. The elongate hemlock scale, Fiorinia externa Ferris (Hemiptera: Diaspididae): A new look at an old nemesis. In: Onken B, Reardon R, Lashomb J editors. Proceedings of the Hemlock Woolly Adelgid in the Eastern United States Symposium. East Brunswick, NJ 5-7 2005 pp. 248-253. Parker BL, Skinner M, Gouli V, Gouli S, Marcelino J, Carlson J, Schartzberg L. 2005. Management of elongate hemlock scale with entomopathogenic fungi. In: Onken B, Reardon R editors. 3rd Symposium on Hemlock Woolly Adelgid in the Eastern United States. Asheville, NC 1-3 February 2005 pp. 161-168. Peres NA, Timmer LW, Adaskaveg JE, Correll JC. 2005. Lifestyles of Colletotrichum acutatum. Plant Disease 89: 784-796. Prusky D, Freeman S, Dickman MB. 2000. Colletotrichum – Host specificity, Pathology and Host-Pathogen Interaction. APS Press. SAS Institute. 1990. SAS/STAT user's guide. 4th ed. Cary, NC. SAS Institute. Sasscer ER. 1912. The genus Fiorinia in the United States. USDA Bur. Entomol. Tech. Ser. 16: 75-82. Smith KT, Lewis PA. 2005. Potential concerns for tree wound response from stem injections. In: Onken B, Reardon R editors. 3rd Symposium on Hemlock Woolly Adelgid in the Eastern United States. Asheville, NC 1-3 February 2005 pp. 173-178. TeBeest DO. 1988. Additions to host range of Colletotrichum gloeosporioides f. sp. aeschynomene. Plant Disease 72: 16-21. Tu JC. 1992. Colletotrichum lindemuthianum on bean: Population dynamics of the pathogen and breeding resistance. In: Bailey JA, Jeger MJ, editors. Colletotrichum: Biology, Pathology and Control, pp. 203-224. CAB International. Zhu P, Oudemans PV. (2000). A retrotransposon Cgret from the phytopathogenic fungus Colletotrichum. Current Genetics 38: 241-247.

143 CHAPTER 5

Entomopathogenic Activity of a Colletotrichum acutatum Strain Recovered from Elongate Hemlock Scale

José A. P. Marcelino1a, Svetlana Gouli1, Vladimir V. Gouli1, Bruce L. Parker1, Margaret Skinner1

1 Entomology Research Laboratory, The University of Vermont (UVM), Burlington, VT 05405-0105 USA

Abstract

A fungal epizootic in populations of Fiorinia externa infesting hemlock trees in forests of the northeastern US has been recently detected. The current known distribution of the epizootic spans 36 sites in New York, Pennsylvania, New Jersey and Connecticut. Colletotrichum acutatum f. sp. fiorinia was the most prevalent fungus recovered from infected scales. Bioassays indicated that this Colletotrichum sp. is highly pathogenic to F. externa. Mortality rates of >90 and >55% were obtained for F. externa crawlers and settlers, respectively. Other insect species from different orders were bioassayed and mortality of ≤22% was obtained suggesting this fungus may be fairly host specific. This is the second report of a Colletotrichum sp. with pathogenic activity towards a scale insect. Our data suggest that C. acutatum f. sp. fiorinia from F. externa epizootics in the US and the previously reported C. gloeosporioides f. sp. ortheziidae from Orthezia praelonga epizootics in Brazil, may constitute distinct biotypes of Colletotrichum that have attained the ability to infect insect hosts in addition to the commonly reported plants.

Keywords: Fiorinia externa, Colletotrichum sp., epizootic, elongate hemlock scale, Orthezia praelonga Corresponding author: a [email protected]

144

Introduction

The eastern hemlock, Tsuga canadensis (L.) Carrière, a common species in forests of the northeastern United States is in decline (Orwig et al. 2002). The invasive elongate hemlock scale, Fiorinia externa Ferris (Hemiptera: Diaspididae), has been identified as one of the causal agents of this decline (Lambdin et al. 2005). Attempts to control this pest have not been successful. The unique shield-like cover of the scale provides protection from contact insecticides, natural enemies and adverse climatic conditions. Because of its high reproductive rate, even when high mortality exceeding

90% occurs, populations quickly rebound (Baranyovits 1953, Johnson and Lyon 1988).

In 2002 an epizootic was reported within the population of F. externa in the

Mianus River Gorge Preserve in Bedford, NY (McClure 2002). Sclerotia were found concealing the bodies of adult mummified scales. The origin of the epizootic was unknown, but similar epizootics have been found among scales in 36 different sites in

New York, Pennsylvania, Connecticut and New Jersey. A complex of entomopathogenic, phytopathogenic and saprophytic fungi were morphologically and molecularly identified as being associated with the diseased insects (Marcelino et al., submitted). One species,

Colletotrichum acutatum f. sp fiorinia (Marcelino et al., submitted) [hereafter referred to

as the Fiorinia strain], was dominant in this complex and was consistently recovered in

F. externa populations in most of the epizootic sites.

Literature on the phytopathogenic genus Colletotrichum (Bailey and Jeger 1992,

Prusky et al. 2000) includes a single report of the species C. gloeosporioides causing

significant epizootics in the scale, Orthezia praelonga Douglas 1891 (Hemiptera:

145 Ortheziidae), a major pest of citrus in Brazil. This fungus, C. gloeosporioides f. sp. ortheziidae is under commercial development for management of O. praelonga. (Cesnik

et al. 1996). The epizootic caused by the Fiorinia strain is the second report of a member

of this genus infecting a scale insect. To better understand the role of this fungus in the F.

externa epizootic, the virulence of the Fiorinia strain to four insect species from three

orders (Hemiptera, Lepidoptera and Thysanoptera) was evaluated.

Materials and Methods

Isolates. The virulence of five Fiorinia strains from different areas of the F.

externa epizootic in the Northeast was tested (Table 5.1). In addition, the following other

multiple-spored isolates were also assayed: C. gloeosporioides f. sp. ortheziidae from the

Brazilian epizootic in O. praelonga, (ARSEF4360) (obtained from the Agricultural

Research Service Entomopathogenic Fungal Collection, Cornell University, Ithaca, NY),

two phytopathogenic C. acutatum, one isolated from blueberry (ERL1379) and one from

tomato (ERL1380) (in permanent storage at the Univ. of VT Entomology Research

Laboratory (UVM ERL) Worldwide Collection of Entomopathogenic Fungi, Burlington,

VT), Lecanicillium lecanii (Zimmerman) Gams & Zare (EHS132), an entomopathogenic

strain isolated from F. externa and one Metarhizium anisopliae (CA-1), recovered from litter in a California avocado orchard (used only in the Frankliniella occidentalis bioassays because of its virulence to this insect). Isolates were grown on potato dextrose agar medium (PDA) (39 g/l) supplemented with penicillin (5 ml/l) and streptomycin

(12.5 ml/l). Conidia were harvested 10-12 d after inoculation.

146 Table 5.1. Fungal isolates tested in the insect bioassays

Fungus Geographic Year of Species Code Host type origin collection

Entomo- Colletotrichum f. sp. EHS41 Fiorinia Mohonk, NY 2005

pathogenic fiorinia (Fiorinia externa

fungi strain)

Fiorinia strain EHS48 F. externa Bayberry Lane, NY 2005

Fiorinia strain EHS51 F. externa Esopus, NY 2005

Fiorinia strain EHS58 F. externa Ward Pound Ridge 2005

Reservation, NY

Fiorinia strain EHS61 F. externa Ward Pound Ridge 2005

Reservation, NY

Lecanicilium lecanii EHS132 F. externa South Salem, NY 2005

Metarhizium CA-1 Soil from Temecula, CA 2001

anisopliaea avocado

plantation

C. gloeosporiodes ARSEF4360 Orthezia Jaguariuna, Sao 1994

sp. f. ortheziidae praelonga Paulo, Brazil

Phyto- C. acutatum ERL1379 Blueberry New Jersey 2005 pathogenic fruit fungi C. acutatum ERL1380 Tomato fruit Burlington, VT 2005

a Used in the F. occidentalis bioassays only.

147 Insects

The virulence of the above fungal isolates was tested against several insect

species representing three orders, Hemiptera, Thysanoptera and Lepidoptera, to better

understand their comparative infectiveness.

Bemisia argentifolii (Hemiptera: Aleroydidae). Silverleaf whitefly was reared on

poinsettia (Euphorbia pulcherrima) at the UVM ERL according to the protocol of Negasi et al. (1998). For these bioassays, terminal leaves of 2-3 week old bean plants, Phaseolus vulgaris (var. Royal Burgundy) were excised and the petiole placed in an Oasis® rooting

cube (Smithers-Oasis USA, Kent, OH) held in place by absorbent cotton wool. Cubes

were placed in tap water in 9 mm diam Petri dishes until roots formed (~4 d), then 18

mating pairs of whiteflies were removed from poinsettia, anesthetized for 2 s with carbon

dioxide and placed on each bean leaf. Infested leaves were held in vented plastic boxes

(8.7 cm wide × 9.5 cm long) at 16:8 LD, 75% RH and 24oC. Adults were removed after

24 h to ensure age homogeneity of the progeny. On each leaf 40-180 1st instars were

produced.

Fiorinia externa (Hemiptera: Diaspididae). Scales were field-collected from

understory eastern hemlock trees at the Mount Tom Forest Preserve, Holyoke, MA,

which is located outside the known area of the F. externa epizootic. One day prior to the

bioassay, 30 branches (50 cm long) with new growth and infested with a healthy

population of F. externa crawlers (i.e., 1st instar mobile nymphal stage emerged from the

3rd instar adult female exuvia) and settlers (i.e., 2nd instar immobile nymphal stage after

inserting stylets in the epidermal cells of hemlock leaves, loosing their legs and

148 remaining anchored for life) were randomly sampled. Branches were kept cool during transport and held at 4oC prior to treatment. Eighty 10 cm long terminal twigs with new

growth were clipped from the branches for the assay. On each infested twig a population

of 10-200 settlers and 10-46 crawlers were counted. Freshly pruned branches were

gathered for each assay repetition.

Frankliniella occidentalis (Thysanoptera: Thripidae). Western flower thrips were

reared at the UVM ERL on bean leaves according to the protocol of Doane et al. (1998).

Two day old 2nd instars were used for testing. Each replicate of the assay had 10

thrips/leaf.

Spodoptera exigua (Lepidoptera: Noctuidae). Beet armyworm eggs were

purchased from Benzon Research, Inc. (Carlisle, PA). Upon delivery, eggs were allowed

to hatch in a glass container (12 cm diam. x 20 cm high) containing cabbage leaves. The

containers were wiped with an antistatic tissue before introduction of eggs. The glass

containers were held for 3-4 d at 22 ± 0.5oC and 16:8 LD. After hatching 20 1st instars were randomly selected for each replicate.

Bioassays

For all bioassays, fungal concentrations of 106 and 107 conidia/ml-1 suspended in

sterile distilled water (SDW) and 0.02% Silwet were tested. A suspension of SDW and

0.02% Silwet was used for the controls in the F. externa trials. SDW served as the

controls for the other test insect species. Each insect bioassay was repeated three times

with four replicates for each treatment.

149 For the whitefly assays, each treatment consisted of four leaves, each with 40-180

2 d old 1st instars. A Potter Precision Laboratory Spray Tower (Burkard Manufacturing

Co. Ltd., Rickmansworth, Hertfordshire, UK) operating at 0.84 kg/cm2 with a 0.25 mm diam nozzle was used to spray 2.5 ml of the fungal suspension. Mortality was assessed after 30 d. Individual insects were first inspected for morphological changes in the cuticle or body, i.e., changes in color or body turgor. In cases where mortality could not be confirmed visually, a fluid sample from the hemolymph was mounted on a glass slide and the cell structures were observed for evidence of disruption or death, as compared with that from live, healthy individuals.

For the F. externa assays, isolates were tested against settlers and crawlers using modified protocols of Rose (1990) and Butt & Goettel (2000). For each treatment, four twigs, each containing ≥10 crawlers or settlers, were held vertically in a metal test tube rack and individually sprayed at a distance of 38 cm with a micro droplet mist of 250 µl of conidial suspension using a hand-held plastic spray bottle. Twigs were allowed to air dry for 2 min, then placed individually in sterile graduated 50 ml conical plastic tubes

(Quikrete, Atlanta, GA) containing 16 g of sterilized sand and 7 ml SDW. The sand served to hold the twigs upright and furnished water to prevent desiccation and maintain a humid environment. Each tube was covered loosely with a cap which allowed ventilation. Tubes were placed in plastic bags to prevent desiccation and held at 22oC with 16:8 LD. Mortality of crawlers and settlers was determined 21 d after treatment as described above for the whitefly assay.

150 For the thrips assays, bean leaf discs (3.3 cm diam) were placed on moist filter

paper in 3.5 cm diam Petri dishes, to which 10-2nd instars were added. Each Petri dish was sprayed with 2 ml of the fungal suspension using a Potter Spray Tower, as described previously. After being air dried for 2 min, Petri dishes were covered and sealed with parafilm, and held in the dark at 22 ± 0.5ºC. Mortality was assessed 7 d after treatment.

Insects that exhibited obvious signs of fungal infection, i.e., displayed an abnormal body color or lacked turgor, and those that did not respond when gently probed with a small insect pin were considered dead.

Beet armyworms were assayed in well plates with 5 x 4 cells (13 mm diam/cell)

(Model #BIO-BA-128, C-D International, Pitman, NJ). A 10 mm diam disc of moist filter

paper was placed in the bottom of each well, followed by 10 mm diam cabbage leaf disc

and one 1st instar beet armyworm. Each 20-cell unit was sprayed with 2 ml of the test

suspension with a Potter Spray Tower. Cell units were air dried for 2 min, covered with

clear plastic wrap and held in an incubator in the dark at 22 ± 0.5°C. Mortality was

assessed after 7 d as described for the thrips assays.

To verify Koch’s postulates (i.e., re-isolation of the test fungi from a diseased host

after treatment of a healthy individual), a random subsample of 10 insects from each

treatment was taken. Each insect was surface sterilized in 0.01% bleach, rinsed in SDW

and placed in a Petri dish on PDA with 5 ml/l penicillin and 12.5 ml/l streptomycin. Petri

dishes were held in the dark at 22 ± 0.5oC for 7 d, and then cadavers were examined for

the presence of the Fiorinia strain or the other test strains.

151 Statistical analyses

Because protocols differed among the insect species bioassayed, statistical

analyses of mortality were made separately for each species. Transformation of the data

was not required as they were normally distributed. Variances were not homogeneous

(using Levene's test), which required use of a Welch’s one way ANOVA (unpooled

variances). An adjusted pairwise comparison between fungal isolates within a test insect

species was made using a post-hoc Tukey-Kramer test. The effect of suspension

concentration (106 or 107 conidia/ml-1) was determined with an adjusted least square

means (LS means). P < 0.05 was considered statistically significant. All statistical

analyses were performed using SAS® (SAS Institute 1990) and plotted using SPSS®

(SPSS Inc. 2005).

Results

Definite symptoms and signs of infection were observed among the insects treated with the Fiorinia strains, demonstrating its entomopathogenic capacity (Figure 5.1).

Mortality varied depending on isolate, conidial concentration and insect species tested

(Table 5.2 and 5.3).

The mean percent mortality among F. externa crawlers and settlers treated with the Fiorinia strains ranged from 78-90% and was significantly greater than mortality for all of the other test isolates and the blank control (P < 0.05) (Figure 5.2). In general mortality from the Fiorinia strains was about 30% greater among the crawlers than the settlers. For the bioassay with crawlers, the mortality obtained from the two other known entomopathogens, the Ortheziidae strain and EHS132, was not significantly different from

152 one of the plant pathogens, ERL1379. Mortality following treatment with the other plant

pathogen, ERL1380, was significantly greater than the two entomopathogenic strains.

These results demonstrate that other C. acutatum strains, known to be plant pathogens

have the potential to infect insects, but their virulence was not consistent within or

between insect species. Differences in mortality between the two conidial concentrations

were not significant for any of the isolates tested against F. externa for crawlers and

settlers.

Figure 5.1. Fungal structures from bioassays of the Fiorinia strain and symptomatic and non- symptomatic F. externa. A) acervuli arising from head of an infected whitefly; B) Crawler with

septicemia from EHS51; C) septicemia in F. externa adult from EHS41; D) generalized septicemia in crawlers from EHS51 treatment; E) arrested development in diseased F. externa; F) normal development of the scale cover in F. externa from controls; G) normal development of the scale cover in nature; H) mycelium arising from adult F. externa.

153 Table 5.2. Results of statistical analysis for bioassays with Fiorinia externa immatures

F. externa settlers F. externa crawlers (N=12,343) (N=989)

F P df F P df Main Effect (Welch’s one way 33.19 <0.0001 9 * 41.72 <0.0001 9 * ANOVA) Test isolates

EHS41 0.75 0.38 1 0.07 0.79 1

EHS48 0.00 0.97 1 0.66 0.41 1

EHS51 0.51 0.47 1 0.00 0.94 1

EHS58 1.12 0.29 1 1.01 0.31 1

EHS61 0.03 0.86 1 0.58 0.44 1 (LS means) ERL1379 1.20 0.27 1 0.61 0.43 1

ERL1380 0.01 0.92 1 1.04 0.30 1 Mortality (Isolate * concentration) EHS132 0.01 0.93 1 0.14 0.70 1

ARSEF4360 0.07 0.79 1 0.18 0.67 1

Blank 0.29 0.59 1 0.00 0.98 1

* Significant differences in percent mortality.

▪ Not tested.

154 Table 5.3. Statistical analysis for bioassays with Spodoptera exigua, Bemisia argentifolii and Frankliniella occidentalis

S. exigua B. argentifolii F. occidentalis (N=1,200) (N=11,492) (N=1,320)

Main Effect F P df F P df F P df (Welch’s one way ANOVA) 3.07 0.0029 9* 3.79 0.0006 8* 1.33 0.24 10 Test

isolates

EHS41 10.91 <0.001 1* 3.87 0.05 1 1.44 0.23 1

EHS48 17.67 <0.001 1* 2.21 <0.13 1 0.00 1.00 1

EHS51 20.28 <0.001 1* 1.19 <0.27 1 0.36 0.54 1

EHS58 0.81 0.368 1 6.49 <0.01 1* 3.24 0.07 1

EHS61 0.36 0.543 1 6.45 <0.01 1* 1.44 0.23 1

ERL1379 1.44 0.231 1 1.56 <0.21 1 1.44 0.23 1 (LS means)

ERL1380 3.24 0.073 1 4.32 <0.03 1* 5.76 0.01 1*

EHS132 70.67 <0.001 1* 7.41 <0.007 1* 0.00 1.00 1 Mortality (Isolate * concentration)

ARSEF4360 0.00 1.00 1 ▪ ▪ ▪ 0.00 1.00 1

CA-1 ▪ ▪ ▪ ▪ ▪ ▪ 81.03 <0.0001 1*

SDW 3.24 0.073 1 0.01 0.93 1 0.36 0.54 1

* Significant differences in percent mortality.

▪ Not tested.

155

Fiorinia externa crawlers Fiorinia externa settlers Fiorinia ex ternasettlers 100 aa a aa 100 I 6 -1 I I 10 conidia/ml I I I I I I 107 conidia/ml-1 I

75 75 a )

) c a a )

) a

% %

% % (

( a

( I (

e e e

es es t

t c

t t

a a

r r

50 50 I I I y

y I I

y y t

t I

t t i

b b i I

i i l

l l

l d a

I b a

a a I I

t t

r r r

r I o

I o o

o I

M M M M I I d I I d 25 25 b I I I

I I I b I

I 0 0 EHS41 EHS51 EHS61 ERL1379ERL1379 EHS132EHS132 EHS41 EHS51 EHS61 ERL1379ERL1379 EHS132EHS132 0 EHS48 EHS58 ARSEF4360ARSEF436 ERL1380 BLANKSDW EHS48 EHS58 ARSEF4360ARSEF4360 ERL1380ERL1380 BLANKSDW

Isolate Isolate

Figure 5.2. Percent mortality among F. externa settlers and crawlers in fungal bioassays. (± 95%

CI of the mean). Bars with the same letters are not significantly different (data for both concentrations combined) (P < 0.05) using the post-hoc multiple comparison Tukey-Kramer test.

Mortality of ≤22% was obtained for all of the other insect species tested (P <

0.05) (Figure 5.3). For some of the test isolates, significantly greater mortality was obtained from the high concentration, though no consistent pattern was observed across insect species or isolates. Of all the insect species tested, whiteflies appeared to be more susceptible to infection at the higher concentration than the other insects. Whiteflies treated with the plant pathogenic strains of C. acutatum exhibited mortality rates comparable with those of the Fiorinia strains.

156

Bemisia argentifolii Spodoptera exigua B. argentifollii Spodoptera exigua 75 75 6 -1 10 conidia/ml 107 conidia/ml-1

) ) ) )

) ) )

50 ) 50

% % % %

( ( (

(

( ( ( (

e e e e

t t t t

a a a a

r r r r

y y y y

t t t t

i i i

i i i i

l l l l

a a a a

t t t t

r r r r

o o o o

M M M M

M M M 25 I I M 25 I I I d I I I aaa I I a I I b b b ● c I b I I I I I I I I I I I I I I I I I I I I I 0 0 I I I EHS41 EHS51 EHS61 ERL1379 EHS132 EHS41 EHS51 EHS61 ERL1379ERL1379 EHS132 0 EHS48 EHS58 ARSEF4360ARSEF436 0 ERL1380 SDW EHS48 EHS58 ARSEF4360ARSEF4360 ERL1380 SDW

Isolate Isolate

FrankliniellaFrankliniella occidentalisoccidentalis 75

) ) )

) 50

% % % %

( ( ( (

e e e e

t t t

a a a a

r r r r

y y y y

t t t t

i i i i

l l l l

a a a a

t t t

r r r r

o o o o

M M M M 25 a

I b b bbb b b b b I b I I I I 0 I I I I I I I I I I I I I I I EHS41 EHS51 EHS61 ERL1379 EHS132 SDW EHS48 EHS58 ARSEF4360ARSEF436 ERL1380 MetMetCA-1 CA-1

Isolate

Figure 5.3. Percent mortality among different test insect species in fungal bioassays. (± 95% CI

of the mean). Bars with the same letters are not significantly different (data for both

concentrations combined) (P < 0.05) using the post-hoc multiple comparison Tukey-Kramer test.

● Statistical significance not estimated for this isolate because data for only one concentration

was collected.

157 Koch’s postulates were successfully achieved for the Fiorinia strains in silverleaf whitefly, F. externa and western flower thrips. In addition, whereas F. externa in the control treatments and those sprayed with C. gloeosporioides f. sp. ortheziidae underwent normal development and reached maturity (Figure 5.1F-G), normal development was halted among F. externa treated with the Fiorinia strains and phytopathogenic C. acutatum. Infected F. externa settlers did not attain maturity (Figure 5.1E) and settlers and crawlers showed symptoms of mycosis (Figure 5.1D-E).

Discussion

This research clearly demonstrates that the Fiorinia strains, isolated originally from F. externa from several sites where a fungal epizootic has been observed, is highly pathogenic to that host, particularly in the crawler stage. In addition Koch’s postulates were satisfied as these test isolates were recovered from cadavers proving that they were killed by infection from these fungi. Infection was also obtained when F. externa crawlers and settlers were treated with other C. acutatum isolates known to be phytopathogenic, though mortality rates were significantly lower than those obtained from the epizootic. In contrast, other insects from the same and different orders displayed comparably lower susceptibility to both the entomopathogenic and phytopathogenic

Colletotrichum isolates.

The present report of the Fiorinia strain and its virulence to F. externa along with the research demonstrating the infectivity of C. gloeosporioides f. sp. ortheziidae to scales of citrus supports the hypothesis that members of the genus Colletotrichum have a broader host range and niche than what has currently been reported (Guerber and Correll

158 2001; Peres et al. 2005). Considering the common occurrence of epizootics linked with

the Fiorinia strain in populations of F. externa within hemlock forests in the Northeast,

this can not be considered an unusual occurrence that takes place under very specific

conditions. Based on the bioassays representing only a few insect species, it appears that this host range extension by the Fiorinia strain is limited primarily to scale insects, or perhaps to Hemiptera in general. Samples of other insects occurring within the epizootic have been sampled, representing a wide variety of species and orders, yet no evidence of infection by the Fiorinia strain in these specimens were obtained (J.M., unpublished

data).

Many questions regarding this unique phenomenon remain that must be answered

before exploitation of this potential biological control agent can be considered. Extensive

sampling within F. externa populations in and outside the epizootic area is needed to

determine the geographical range of the Fiorinia strain. This would provide essential

information on its current distribution, its rate of expansion and dispersal, and the site of

origin. Currently it is unknown if this entomopathogenic strain occurred here before the

accidental introduction of F. externa or arrived in association with the scale. It is possible

that in its native habitat, this fungus maintains it naturally at low levels. The process by

which infection of this fungus takes place also requires further study. The Fiorinia strain

has been found growing endophytically in over 28 different species of plants within the

epizootic areas, though external evidence of infection (signs and symptoms) is lacking

(J.M., unpublished data). It is unknown if F. externa becomes infected through contact

with these plants or some other means. This research shows many aspects of fungal

159 ecology are only now being discovered, even for organisms such as Colletotrichum which has been studied extensively for over a century.

Acknowledgments

We thank Tom Doubleday, Cheryl Frank, Teri Hata and Dr. Robert Jones for their help with this project. This work was funded in part through a grant awarded by the

Northeastern Area State and Private Forestry, USDA Forest Service (#04-CA-

11244225286) and is in partial fulfillment of requirements for the PhD degree of J.M. at the University of Vermont.

160 Literature Cited

Baranyovits F. 1953. Some aspects of the biology of armored scales. Endeavour 12: 202- 209. Bailey JA, Jeger MJ. 1992. Colletotrichum: Biology, Pathology and Control. CAB International. 402 p. Butt TM, Goettel MS. 2000. Bioassays of entomogenous fungi. In: Navon A, Ascher KRS editors. Bioassays of Entomopathogenic Microbes and Nematodes, pp.141-195. CABI International Press. Cesnik R, Ferraz JNG, Oliveira RCAL, Arellano F, Maia AH. 1996. Controle de Orthezia praelonga com o fungo Colletotrichum gloeosporioides isolado orthezia, na regiao de Limeira, SP. In: 5o Simpósio de Controle Biológico. Foz de Iguaçu, Brazil, 5-10 June 1996 p. 363 (In Portuguese) Doane EN, Parker BL, LaRosa S, Skinner M, Boone J, Pivot, Y. 1998. Mass rearing of western flower thrips, Frankliniella occidentalis (Thysanoptera: Thripidae) on beans. Agricultural Experimental Station, University of Vermont. Technical Note 4. 6 p. Guerber JC, Correll JC. 2001. Morphological description of Glomerella acutata, the teleomorph of Colletotrichum acutatum. Mycologia 93: 216-229. nd Johnson WT, Lyon HH. 1988. Insects that feed on trees and shrubs. 2 Ed. Cornell Univ. Press. 556 p. Lambdin P, Lynch C, Grant J, Reardon R, Onken B, Rhea R. 2005. Elongate hemlock scale and its natural enemies in the southern Appalachians. In: Onken B, Reardon R editors. 3rd Symposium on Hemlock Woolly Adelgid in the Eastern United States. Asheville, NC 1-3 February 2005 pp. 145-154. McClure MS. 2002. The elongate hemlock scale, Fiorinia externa Ferris (Hemiptera: Diaspididae): A new look at an old nemesis. In: Onken B, Reardon R, Lashomb J editors. Proceedings of the Hemlock Woolly Adelgid in the Eastern United States Symposium. East Brunswick, NJ 5-7 2005 pp. 248-253. Negasi A, Parker BL, Brownbridge M. 1998. Screening and bioassay of entomopathogenic fungi for the control of silverleaf whitefly, Bemisia argentifolii. Insect Science and its Applications 18: 37-44. Orwig DA, Foster DR, Mausel DL. 2002. Landscape patterns of hemlock decline in New England due to the introduced hemlock woolly adelgid. Journal of Biogeography 29: 1475-1487. Peres NA, Timmer LW, Adaskaveg JE, Correll JC. 2005. Lifestyles of Colletotrichum acutatum. Plant Disease 89: 784-796. Prusky D, Freeman S, Dickman MB. 2000. Colletotrichum – Host specificity, Pathology and Host-Pathogen Interaction. APS Press. Rose M. 1990. Rearing and mass rearing. In: Rosen D editor, Armored scale insects. Their Biology, Natural Enemies and Control. [Series title: World crop pests, Vol. 4A: The Armored Scale Insects], pp. 357-365. Elsevier. SAS Institute. 1990. SAS/STAT user's guide. 4th ed. Cary, NC. SAS Institute. 1069 p. SPSS Inc. 2005. SPSS Base 14.0 for Windows User's Guide. Chicago, IL. SPSS Inc. 783 p.

161 CHAPTER 6

Comprehensive bibliography

ADASKAVEG, J.E., AND R.J. HARTIN. 1997. Characterization of Colletotrichum acutatum isolates causing anthracnose of almond and peach in California. Phytopathology 87: 979-987. AFANADOR-KAFURI, L., D. MINZ, M. MAYMON, AND S. FREEMAN. 2003. Characterization of Colletotrichum isolates from tamarillo, passiflora, and mango in Colombia and identification of a unique species from the genus. Phytopathology 93: 579–587. ALEXANDER, S. A., AND J.K. MARVEL. 2001. Colletotrichum gloeosporioides causes severe anthracnose disease on pepper in Virginia. P. 1 in Joint meeting of the northeastern and potomac divisions. American Phytopathological Society. Cromwell, Connecticut. (Abstr.) ALTSCHUL, S.F., W. GISH, W. MILLER, E.W. MYERS, AND D.J. LIPMAN. 1990. Basic local alignment search tool. J. Mol. Biol. 215: 403–410. ANAGNOSTAKIS, S.L. 1988. Cryphonectria parasitica: cause of Chestnut blight. Advances in Plant Pathology 6: 123-136. ANDERSON, H.W., AND A.G. GORDON.1994. The tolerant conifers: Eastern hemlock and red spruce, their ecology and management. Ontario forest research institute. Forest Research Report 113. 114 p. ANSARI, K.I., N. PALACIOS, C. ARAYA, T. LANGIN, D. EGAN, AND F.M. DOOHAN. 2004. Pathogenic and genetic variability among Colletotrichum lindemuthianum isolates of different geographic origins. Plant Pathology 53: 635-642. ANNECKE, D.P. 1963. Observations on some citrus pests in Moçambique and southern Rhodesia. Journal Ent. Soc. S. Africa 26 (1): 194-225. ARROYO, F.T, J. MORENO, G. GARCÍA-HERDUGO, B. DE LOS SANTOS, C. BARRAU, M. PORRAS, C. BLANCO, AND F. ROMERO. 2005. Ultrastructure of the early stages of Colletotrichum acutatum infection of strawberry tissues. Can. J. Bot. 83: 491-500. AZEVEDO, J.L., W. MACCHERONI, J.O. PEREIRA, AND W.L ARAUJO. 2000. Endophytic microorganisms: a review on insect control and recent advances on tropical plants. Electronic J. of Biotechnology 3(1): 40-61. BARANYOVITS, F. 1953. Some aspects of the biology of armoured scales. Endeavour 12: 202-209. BAILEY, J.A., R.J. O’CONNELL, R.J. PRING, AND C. NASH. 1992. Infection strategies of Colletotrichum species. P. 88-120 in Colletotrichum: Biology, pathology and control, Bailey, J. A. and M. J. Jeger (eds.). CAB International. Wallingford, UK. BAILEY, J.A., AND M.J. JEGER. 1992. Colletotrichum: Biology, pathology and control. CAB International. Wallingford, UK. 400 p. BAILEY, J.A., C. NASH, L.W. MORGAN, R.J. O’CONNELL, AND D.O. TEBEEST. 1996. Molecular taxonomy of Colletotrichum species causing anthracnose on the Malvaceae. Phytopathology 86: 1076-1083.

162 BALARDIN, R.S., J.J. SMITH, AND J.D. KELLY. 1999. Ribosomal DNA polymorphism in Colletotrichum lindemuthianum. Mycol. Res. 103 (7): 841-848. BALDWIN, J.L. 1973. Climates of the United States. U. S. Dep. Commerce, Washington, D. C. 113 p. BARNETT, H.L., AND B.B. HUNTER. 1998. Illustrated genera of imperfect fungi 4th edition. APS Press, St. Paul, MN. 218 p. BARTA, M., AND L. CAGAN. 2003. Entomophthoralean fungi associated with the common nettle aphid (Microlophium carnosum Buckton) and the potential role of nettle patches as reservoirs for the pathogens in landscape. J. Pest Sci. 76 (1):6–13. BATISTA, A.C., AND J.L. BEZERRA. 1965. Sobre o parasitismo do Colletotrichum gloeosporioides Penz. e outros fungos, em Orthezia praelonga Douglas. Brotéria 35: 68-74. (In Portuguese, abstr. in English) BAXTER, A.P., G.C.A. VAN DER WESTHUIZEN, AND A. EICKER. 1983. Morphology and taxonomy of South African isolates of Colletotrichum. S. African J. of Botany 2 (4): 259-289. BAXTER, A.P., G.C.A. VAN DER WESTHUIZEN, AND A. EICHER. 1985. A review of literature on the taxonomy, morphology and biology of the fungal genus Colletotrichum. Phytophylactica 17: 15-18. BEARDSLEY, J.W. 1969. A new fossil scale insect from Canadian amber. Psyche 76: 270- 279. BEARDSLEY, J.W., AND R.H. GONZALEZ. 1975. The biology and ecology of armored scales. Annu. Rev. Entomol. 20: 47-73. BECHINGER, C., K.-F. GIEBEL, M. SHNELL, H.B. DEISING, AND M. BASTMEYER. 1999. Optical measurements of invasive forces exerted by apressoria of a plant pathogenic fungus. Science 285: 1896-1899. BENOIT, J.B., J.A. YODER, L.W. ZETTLER, AND H.H. HOBBS. 2004. Mycoflora of a trogloxenic cave cricket, Hadenoecus cumberlandicus (Orthoptera: Rhaphidophoridae), from small caves in northeastern Kentucky. Annals of the Entomological Society of America 97 (3): 989-993. BERBEE, M.L., AND J.W. TAYLOR. 2001. Fungal molecular evolution: Gene trees and geologic time. P. 229-245 in The Mycota - a Comprehensive treatise on fungi as experimental systems for basic and applied research, VII: Systematics and evolution, Part B, Esser K. and J.W. Bennett (eds.). Berlin: Springer-Verlag. BERGER, E.W. 1919. How to introduce the red and yellow whitefly fungi (red Aschersonia and yellow Aschersonia). P. 160-170 in Work of the Entomological Department State Plant Board Fla. St. Hor. Soc., Proc. 32. BERGER, E.W. 1921. Natural enemies of scale insects and whiteflies in Florida. Quart. Bull. Fla. Stn. Plant Board 5: 141. BERGSTROM, G.C., AND R.L. NICHOLSON 2000. The Biology of Colletotrichum graminicola and maize anthracnose. P. 374-393 in Colletotrichum – Host specificity, pathology and host-pathogen interaction, Prusky, D., S. Freeman and M.B. (eds.). APS Press. St. Paul, Minnesota. BENJAMIN, R.K., M. BLACKWELL, I.H. CHAPELA, R.A. HUMBER, K.G. JONES, K.D KLEPZIG, R.W. LICHTWARDT, D. MALLOCH, H. NODA, R.A. ROEPER, J. SPATAFORA,

163 AND A. WEIR. 2004. Insect – and other arthropod – associated fungi in Biodiversity of fungi – Inventory and monitoring methods, Muller, G., M. Foster and G. Bills (eds.). Elsevier. 777 p. BISIACH, M. 1992. Convegno sulla lotta biologica al cancro corticale del castagno. Centro studi per il nocciolo ed il castagno, Caprarola (Viterbo): 10-21. (In Italian). BLUMENTHAL, E.M. 2003. Biological control of elongate hemlock scale. FPM News, Vol. 21 (2): 5-6. BOONPHONG, S., P. KITTAKOOP, M. ISAKA, P. PALITTAPONGARNPIM, A. JATURAPAT, K. DANWISETKANJANA, M TANTICHAROEN, AND THEBTARANONTH, Y. 2001. A new antimycobacterial, 3β-Acetoxy-15α, 22-dihydroxyhopane, from the insect pathogenic fungus Aschersonia tubulata. Planta Medica 67: 279-281. BONNEAU, L.R., K.S. SHIELDS, AND D.L CIVCO. 1999. Using satellite images to classify and analyze the health of hemlock forests infested by the hemlock woolly adelgid. Biological Invasions 1: 255-267. BORISOV, B.A., V.V. SEREBOV, I.I. NOVIKOVA, AND I.V. BOYKOVA. 2001. Insect pathogens: Structural and functional aspects. P. 352-427 in Entomopathogenic fungi: Ascomycota and Deuteromycota. BOUCIAS, D.G., AND J.C. PENDLAND 1983. Host recognition and specificity of entomopathogenic fungi. P. 185-196 in Infection processes of fungi, Roberts, D. W. and J. R. Aist (eds.). Rockefeller Fdn., NY. ® BOWERS, R.C. 1986. Commercialization of Collego - An industrialist’s view. Weed Sci. 34: S24-S25. BRAY, D. F. 1958. Fiorinia hemlock scale. Scientific Tree Topics 2(5): 11. BROOKER, N.L., J.F. LESLIE, AND M. B. DICKMAN. 1991. Nitrate non-utilizing mutants of Colletotrichum and their use in studies of vegetative compatibility and genetic relatedness. Phytopathology 81: 672-677. BROOKS, A.N. 1931. Anthracnose of strawberry caused by Colletotrichum fragariae, n. sp. Phytopathology 21: 739-744. BROWNING, M., L.V. ROWLEY, P. ZENG, J.M. CHANDLEE, AND N. JACKSON. 1999. Morphological, pathogenic, and genetic comparisons of Colletotrichum graminicola isolates from poaceae. Plant Disease 83 (3): 286-292. BRYSON, R.J., C.E. CATEN, D.W. HOLLOMON, AND J.A. BAILEY. 1992. Sexuality and genetics of Colletotrichum. P. 27-46 in Colletotrichum – Biology, Pathology and Control, Bailey, J.A. and M.J. Jeger (eds.). CAB International. Wallingford, UK. BURNS, G.P. 1923. Studies in tolerance of New England forest trees. V. Minimum light requirements referred to a definite standard. Vermont Agricultural Experiment Station Bulletin 235: 1-31. BURGES, H.D. 1981. Microbial control of pests and plant diseases 1970-1980. Academic Press, New York. 949 p. BURGES, H.D., AND HALL, R.A. 1982. Bacteria and fungi as insecticides. Outlook on Agriculture 11 (2). 79 p. BUTT, T.M, AND M.S. GOETTEL. 2000. Bioassays of entomogenous fungi. P.141-195 in Bioassays of entomopathogenic microbes and nematodes, Navon, A. and K.R.S. Ascher (eds.). CABI International Press, Wallingford, U.K.

164 BYRNE, J.M., M.K. HAUSBECK, AND R. HAMMERSCHMIDT. 1997. Conidial germination and apressorium formation of Colletotrichum coccodes on tomato foliage. Plant Disease 81: 715-718. CAMPBELL, R.K., T.E. ANDERSON, M. SEMEL, AND D.W. ROBERTS. 1985. Management of Colorado potato beetle using entomogenous fungus Beauvaria bassiana. American Potato Journal 62: 29-37. CANADIAN FOOD INSPECTION AGENCY. 1997. Deregulation of Colletotrichum acutatum, Simmonds. Animal and Plant Health Directorate. Plant Protection Division. File 3525-11F1-0/3525-11F1/FN7 3510-3-1B1. http://www.inspection.gc.ca/english/plaveg/protect/dir/d-97-03e.pdf (Last access in 06/04/2005) CANNON, P.F., P.D. BRIDGE, AND E. MONTE. 2000. Linking the past, present and future of Colletotrichum systematics. P. 1-20 in Colletotrichum – Host specificity, pathology and host-pathogen interaction, Prusky, D., S. Freeman and M.B. Dickman (eds.). APS Press. St. Paul, Minnesota. CANO, J., J. GUARRO, AND J. GENÉ. 2004. Molecular and morphological identification of Colletotrichum species of clinical interest. J. of Clinical Microbiol. 42 (6): 2450- 2454. th CARLISLE, M.J., AND S.C. WATKINSON. 1997. The fungi. 4 Edition. Academic Press. Boston, Massachusetts, USA. CARMICHAEL, J.W., W.B. KENDRICK, I.L. CONNERS, AND L. SIGLER. 1980. Genera of Hyphomycetes. The University of Alberta Press. 369 p. CARROL, G. 1995. Forest endophytes: pattern and process. Can. J. Bot. 73: S1316-S1324. CARSON, R. 1987. Silent Spring Revisited. Edited by Washington DC: American Chemical Society. 214 p. CASTRO, V.L., R. CESNIK, AND R.C.A.L OLIVEIRA. 1998. Testes toxicológicos em ratos expostos ao Colletotrichum gloeosporioides. P. 156 in 6o Simpósio de Controle Biológico. Rio de Janeiro, Brazil. (In Portuguese) CERKAUKAS, R.F. 1988. Latent colonization by Colletotrichum spp.: epidemiological considerations and implications for mycoherbicides. Can. J. Plant Pathol. 10: 297- 310. CESNIK, R., AND G.C.G. OLIVEIRA. 1993. Colletotrichum gloeosporioides isolado de Orthezia praelonga causando patogenicidade em Coccoloba sp. Summa phytopathologica 19 (1): 37. (In Portuguese) CESNIK, R., J.N.G. FERRAZ, R.C.A.L OLIVEIRA, F. ARELLANO, AND A.H. MAIA. 1996. Controle de Orthezia praelonga com o fungo Colletotrichum gloeosporioides isolado orthezia, na regiao de Limeira, SP. P. 363 in: 5o Simpósio de Controle Biológico. Foz de Iguaçu, Brazil. (In Portuguese) CESNIK, R., AND J.N.G. FERRAZ. 2000. Orthezia praelonga, 1891 (Homoptera, Ortheziidae) biologia, controle quimico e biologico. Embrapa Meio Ambiente, Jaguariúna, Brazil, Boletim de Pesquisa 9. 27 p. (in Portuguese) CESNIK, R., H.S. PRATES, AND J.N.G. FERRAZ. 2005. Utilização de fungos entomopatogênicos no controle de Orthezia praelonga na citricultura. Embrapa Meio

165 Ambiente; Ministério da agricultura, pecuária e abastecimento and Governo do estado de São Paulo. Leaflet. (In Portuguese) CHACKO, R.J., G.J. WEIDEMANN, AND D.O. TEBEEST. 1989. Heterokaryosis among subspecies of Colletotrichum gloeosporioides. Phytopathology 79 (10): 1208. (Abstr.) CHACKO, R.J., J.C. CORRELL, G.J. WEIDEMANN, AND D.O. TEBEEST. 1990. Vegetative compatibility among isolates of Colletotrichum gloeosporioides f. sp. aeschynomene. Phytopathology 80 (10): 1057. CHARNLEY, A.K. 1997. Entomopathogenic fungi and their role in pest control. P. 185-201 in The Mycota - a Comprehensive treatise on fungi as experimental systems for basic and applied research, IV: Environmental and microbial relationships. Esser K. and J.W. Bennett (eds.). Berlin: Springer-Verlag. CHARUDATTAN, R. 1991. The mycoherbicide approach with plant pathogens. P. 3-23 in Microbial Control of Weeds, TeBeest, D. O. (ed.). Routledge, Chapman & Hall, Inc. New York. 280 p. CHENNA R., H. SUGAWARA, T. KOIKE, R. LOPEZ, T.J.GIBSON, D.G. HIGGINS, AND J.D. THOMPSON. 2003. Multiple sequence alignment with the Clustal series of programs. Nucleic Acids Res. 31 (13): 3497-500 CHILTON, S.J.P., G.B. LUCAS, AND C.W. EDGERTON. 1944a. Genetics of Glomerella. I. Studies on the behavior of certain strains. Am. J. Bot. 31 (4): 233-239. CHILTON, S.J.P., G.B. LUCAS, AND C. W. EDGERTON. 1944b. Genetics of Glomerella. III. Crosses with a conidial strain. Am. J. Bot. 32 (9): 549-554. CHILTON, S.J.P., AND H.E. WHEELER. 1949a. Genetics of Glomerella. VI. Linkage. Am. J. Bot. 36 (3): 270-273. CHILTON, S.J.P., AND H.E. WHEELER. 1949b. Genetics of Glomerella. VII. Mutation and segregation in plus cultures. Am. J. Bot. 36 (10): 717-721. CHARLES, V.K. 1941. A preliminary check list of the entomogenous fungi of North America. U.S. Bur. Plant. Indus. Insect Survey Bull. 21: 770-785. CISAR, C.R., F.W., SPIEGEL, D.O. TEBEEST, AND C. TROU. 1994. Evidence for mating between isolates of Colletotrichum gloeosporioides with different host specificities. Curr. Gen. 25: 330-335. CISAR, C.R., AND D.O. TEBEEST. 1995. Sexual cycle and potential for gene flow in fungal biological control agents. http://www.isb.vt.edu/brarg/brasym95/cisar95.htm (Last access 8/01/2007) CISAR, C.R., A.B. THORNTON, AND D.O. TEBEEST. 1996. Isolates of Colletotrichum gloeosporioides (Telemorph: Glomerella cingulata) with different host specificities mate on northern Jointvetch. Biological Control 7: 75-83. CLAMP M., J. CUFF, S.M. SEARLE, AND G.J. BARTON. 2004. The Jalview Java alignment editor. Bioinformatics 20: 426-427. COLBERT, J., M. SEESE, AND B. ONKEN. 2002. Hemlock woolly adelgid impact assessment survey. P. 323-328 in Proceedings of the hemlock woolly adelgid in the eastern United States Symposium, Onken, R. R. and J. Lashomb (eds.). New Jersey Agricultural Experiment Station, Rutgers University, New Brunswick, NJ. COLEY-SMITH, J.R., AND R.C. COOKE. 1971. Survival and germination of fungal sclerotia. Annu. Rev. Phytopathol. 9: 65-92.

166 CONNICK, W.J., D.J. DAIGLE, C.D. BOYETTE, K.S. WILLIAMS, B.T VINYARD, AND P.C. QUINBY, 1996. Water activity and other factors that affect the viability of Colletotrichum truncatum conidia in wheat flour-kaolin granules (‘Pesta’). Biocontrol Science and Technology 6: 277-284. COUTO, E.F., AND M. MENEZES. 2004. Caracterização fisiomorfológica de isolados de Colletotrichum musae. Fitopatol. Bras. 29 (4): 406-412. (In Portuguese) CORDA, A.K.J. 1831. Die pilze Deutschlands, in Deutschlands Flora, 3. Abtheilung. Sturm, J. eds. 3 (12): 33-64. (In German). COWLES, R.S., COWLES, E.A., MCDERMOTT, A.M., RAMOUTAR, D. 2000. “Inert” formulation ingredients with activity: toxicity of trisiloxane surfactant solutions to twospotted spider mites (Acari: Tetranychidae). J. Econ. Entomol. 93: 180-88. CROUCH J.A., B.B. CLARKE, AND B.I. HILLMAN. 2005. Diversity in natural populations of Colletotrichum from the tallgrass prairie. P. 61 in 23rd Fungal genetics conference. Von den Hondel, C. and G. Turgeon (eds.). Asilomar, California. (Abstr.) CROUCH, J.A., B.B CLARKE, AND B.I. HILLMAN. 2006. Unraveling evolutionary relationships among the divergent lineages of Colletotrichum causing anthracnose disease in turfgrass and corn. Phytopathology 96: 46-60. CROW, T.R. 1995. The social, economic, and ecological significance of hemlock in the Lake states. P. 11-17 in Hemlock ecology and management. Proceedings of a regional eonference on ecology and management of eastern hemlock. Mroz, G. and A. J. Martin (eds.). Dept. of Forestry, University of Wisconsin, Madison. CURTIS, J.T. 1959. The vegetation of Wisconsin. The Univ. of Wisconsin Press. 645 p. DANIEL, J.T., G.E. TEMPLETON, R.J. JR. SMITH, AND W.T. FOX. 1973. Biological control of northern jointvetch in rice with an endemic fungal disease. Weed Science 21 (4): 303-307. DANOFF-BURG, J.A., AND S. BIRD. 2002. Hemlock woolly adelgid and elongate hemlock scale: Partners in crime? P. 254-268 in Proceedings of the hemlock woolly adelgid in the eastern United States Symposium. Onken, B., R. Reardon, and J. Lashomb (eds.). New Jersey Agricultural Experiment Station, Rutgers University, New Brunswick, NJ. DAOUST, R.A., AND D.W. ROBERTS. 1982. Virulence of natural and insect-passaged strains of Metarhizium anisopliae to mosquito larvae. Journal of Invertebrate pathology 40: 107-117. DAOUST, R.A., AND D.W. ROBERTS. 1983. Studies on the prolonged storage of Metarhizium anisopliae conidia: effect of growth substrate on conidial survival and virulence against mosquitoes. Journal of Invertebrate pathology. 41: 161-170. DAS, G.M., AND R.N. GANGULI. 1961. Coccoids on tea in North-East India. Indian Journal of Entomology 23: 245-256. DAS, E.C., AND S.C. DAS. 1962. On the biology of Fiorinia theae Green (Coccidea: Diaspididae) occurring on tea in north-east India. Indian Journal of Entomology 24 (1): 27-35. DAUCH, A.L., A.K. WATSON, AND S.H. JABAJI-HARE. 2002. Detection of the mycoherbicide Velgo® in velvetleaf field soil using strain-specific primers in The

167 Canadian phytopathological society annual meeting, Waterton Lakes National Park, Alberta, Canada. Can J. Plant Pathol. 24: 383-384. DAVIDSON, J.A., AND C. MCCOMB. 1958. Notes of the biology and control of Fiorinia externa Ferris. J. Econ. Entomol. 51: 405-406. DANZIG, E.M. 1961. Food forms of Eulecanium franconicum (Lndgr.) (Homoptera: Coccoidea). Entomol. Rev. 40 (3): 310-313. DEAN, R.A. 1997. Signal pathway and apressorium morphogenesis. Ann. Rev. Phytopathol. 35: 211-234. DEMARSAY, A., AND P.V. OUDEMANS. 2001. Reservoirs of Colletotrichum acutatum in dormant and growing highbush blueberry. Phytopathology 92: S143. (Abstr.). DEVNATH, S. 1986. A new scale insect Asterolecanium pustulans (Asterolecaniidae: Coccoidea) recorded on tea in Darjeeling. Two and a Bud. Vol. 33: 27-28. DICKMAN, M.B., T.L BUHR, V. WARWAR, G.M. TRUESDELL, AND C. HUANG. 1995. Molecular signals during the early stages of alfalfa anthracnose. Can. J. Bot. 73: 1169-1177. DICKMAN, M.B. 1998. Signal exchange during Colletotrichum trifolii-Alfalfa interactions, in Workshop on host specificity, pathology and host–pathogen interaction of Colletotrichum. Ramat Rachel, Jerusalem, Israel. Phytoparasitica 26 (4): 9 (Abstr.) DICKMAN, M.B. 2000. Signal exchange during Colletotrichum trifolii-Alfalfa Interactions. P, 205-231 in Colletotrichum – Host specificity, pathology and host- pathogen interaction, Prusky, D., S. Freeman and M. B. Dickman (eds.). APS Press. St. Paul, Minnesota. DICKMAN, M.B. 2004. Apressorium development in Colletotrichum trifolii. Inoculum 55 (4): 13 (Abstr.). DICKSON, R.C. 1951. Construction of the scale covering of Aonidiella aurantii (Mask.). Ann. Ent. Soc. Am. 44: 596-602. DILLARD, H.R., AND A.C. COBB. 1998. Survival of Colletotrichum coccodes in infected tomato tissue and in soil. Plant Disease 82: 235-238. DINGLEY, J.M., AND J.W. GILMOUR. 1977. Colletotrichum acutatum: Simmds. f. sp. pinea associated with “Terminal crook” disease of Pinus spp. New Zealand J. For. Sci. 2 (2): 192-201. DISSELKAMP, C. 1954. The scale formation on the San Jose Scale (Quadraspidiotus perniciosus Comst. 1881). Höfchen-Briefe 7 (3): 105-141. DITOMMASO, A., AND A.K. WATSON. 1995. Impact of a fungal pathogen, Colletotrichum coccodes on growth and competitive ability of Abutilon theophrasti. New Phytol. 131: 51-60. DITOMMASO, A., A.K. WATSON, AND S.G. HALLETT. 1996. Infection by the fungal Pathogen Colletotrichum coccodes Affects Velvetleaf (Abutilon theophrasti)– Soybean competition in the field. Weed Science 44: 924-933. DOANE, E.N., B.L. PARKER, S. LAROSA, M. SKINNER, J. BOONE, AND Y. PIVOT. 1998. Mass rearing of western flower thrips, Frankliniella occidentalis (Thysanoptera: Thripidae) on beans. Agricultural Experimental Station, University of Vermont. Technical Note 4. 6 p.

168 DODD, J.C., A. ESTRADA, AND M.J. JEGER. 1992. Epidemiology of Colletotrichum gloeosporioides in the tropics. P. 308-325 in: Colletotrichum – Biology, pathology and control, Bailey, J.A. and M.J. Jeger (eds.). CAB International. Wallingford, UK. DRORI, N., H. HAIMOVICH-KRAMER, J. ROLLINS, A. DINOOR, Y. OKON, O. PINES, AND D. PRUSLY. 2003. External pH and nitrogen source affect secretion of pectate lyase by Colletotrichum gloeosporioides. Appl. and Environ. Microbiology 69 (6): 3258-3262. DU, M., C.L. SCHARDL, E.M. NUCKLES, AND L.J. VAILLANCOURT. 2005. Using mating- type gene sequences for improved phylogenetic resolution of Colletotrichum species complexes. Mycologia 97: 641-658. DUDA, E.J. 1957. Fiorinia externa Ferris - a scale on hemlock. Sci. Tree Topics. 2 (4): 9- 10. DUMAS B., C. BOREL, C. HERBERT, J. MAURY, C. JACQUET, R. BALSSE, AND M.T. ESQUERRÉ-TUGAYÉ. 2001. Molecular characterization of CLPT1, a SEC4-like RAB/GTPase of the phytopathogenic fungus Colletotrichum lindemuthianum which is regulated by the carbon source. Gene 272: 219-225. DWYER, G., J.S. ELKINTON, AND A.E. HAJEK. 1998. Spatial scale and the spread of a fungal pathogen of gypsy moth. The American Naturalist 152 (3): 485-494. EASTBURN, D.M., AND W.D. GUBLER. 1990. Strawberry anthracnose: Detection and survival of Colletotrichum acutatum in soil. Plant Disease 74: 161-163. EDGERTON, C.W., S.J.P. CHILTON, AND G.B. LUCAS. 1945. Genetics of Glomerella. II. Fertilization between strains. Am. J. Bot. 32 (3): 115-118. EMMETT, R.W., AND D.G. PARBERY. 1975. Appressoria. Ann. Rev. Phytopathol. 13: 147- 165. EVANS, H.C. 1989. Mycopathogens of insects of epigeal and aerial habitats. P. 205-238 in Insect-Fungal interactions Wilding, N., N.M. Collins, P.M. Hammond and J.F. Webber (eds.). Academic Press. London. EVANS, H.C., AND N.L. HYWEL-JONES. 1997. Entomopathogenic fungi. P. 3-27 in Soft scale insects - Their biology, natural enemies and control Vol. 7B, Ben-Dov, Y. and C. J. Hodgson (eds.). Elsevier. Amsterdam, The Netherlands. EVERETT, K.R. 2003. The effect of low temperatures on Colletotrichum acutatum and Colletotrichum gloeosporioides causing body rots of avocados in New Zealand. Australasian Plant Pathology 32: 441-448. EVLACHOVA, A.A. 1974. Entomopathogenic Fungi. Leningrad. USSR Academy of Sciences. 260 p. (In Russian) FAGBOLA, O., AND M.M. ABANG. 2004. Colletotrichum circinans and Colletotrichum coccodes can be distinguished by DGGE analysis of PCR-amplified 18S rDNA fragments. African J. of Biotechnology 3 (3): 195-198. FARGES, J. 1984. Adhesion of the fungal spore to the insect cuticle in relation to pathogenicity. P. 90-110 in Infection processes of fungi, Roberts, D.W. and J.R. Aist (eds.). Publs. Rockefeller Fdn., NY. FAWCETT, H.S. 1948. Biological control of citrus insects by parasitic fungi and bacteria. P. 627-664 in The Citrus Industry, Vol. 2, Batchelor, L. D. and H. J. Webber (eds.), University of California Press, Berkeley, CA.

169 FARGES, J., AND P-H. ROBERT. 1985. Persistance des conidiospores des hyphomycètes entomopathogens Beauvaria bassiana (Bals.) Vuill. et Metarhizium anisopliae (Metsch) Sor. et Paecilomyces fumosoroseus Wize dans le sol, en conditions contrôlées. Agronomie 5 (1): 73-80. (In French) FAWCETT, H.S. 1908a. Fungi parasitic upon Aleyrodes citri. Univ. Fla. Exp. Stud. 1: 1- 41. FAWCETT, H.S. 1908b. Fungi parasitic upon Aleyrodes citri. Thesis for the degree of Master of Science. The Univ. of the State of Florida, 41 p. FAWCETT, H.S. 1944. Fungus and bacterial diseases of insects as factors in biological control. Bot. Rev. 10: 327-348. FAWCETT, H.S. 1948. Biological control of citrus insects by parasitic fungi and bacteria. P. 627-664 in The Citrus Industry, Vol. 2, Batchelor, L. D. and H. J. Webber (eds.), University of California Press, Berkeley, CA. FENG, K.C., B.L. LIU, AND Y.M. TZENG. 2000. Verticillium lecanii spore production in solid-state and liquid-state fermentations. Bioprocesses Biosyst. Eng. 23: 25-29. FERNANDO, T.H.P.S., C.K. JAYASINGHE, AND R.L.C WIJESUNDERA. 2000. Factors affecting spore production, germination and viability of Colletotrichum acutatum isolates from Hevea brasiliensis. Mycol. Res. 104 (6): 681-685. FERRIS, G.F. 1942. Atlas of the scale insects of North America. Vol. IV, Stanford Univ. Press. FISHER, F.E., W.L. THOMPSON, AND J.T. GRIFFITHS. 1949. Progress report on the fungus diseases of scale insects attacking citrus in Florida. The Florida Entomologist 32: 1- 11. FORD, R., S. BANNIZA, W. PHOTITA, AND P.W.J. TAYLOR. 2004. Morphological and molecular discrimination of Colletotrichum truncatum causing anthracnose on lentil in Canada. Australasian Plant Pathology 33: 559-569. FÖRSTER, H., AND J.E. ADASKAVEG, 1999. Identification of subpopulations of Colletotrichum acutatum and epidemiology of elmond anthracnose in California. Phytopathology 89: 1056-1065. FOSTER, D.R., AND T.M ZEBRYK. 1993. Long-term vegetation dynamics and disturbance history of a Tsuga-dominated forest in New England. Ecology 74 (4): 982-998. FOSTER, D.R. 2000. Hemlock’s future in the context of its history: An ecological perspective. P. 1-4 in Proceedings: Symposium on sustainable management of Hemlock ecosystems in eastern North America. McManus, K.A., K. S. Shields, and D. R Souto, (eds.). USDA Forest Service, Northeastern Research Station, Durham, NH. FOSTER, D.R., AND J.F. O’KEEFE. 2000. New England forests through time. Insights from the Harvard forest dioramas. Harvard Univ. Press. 67 p. FOWELLS, H.A. 1965. Eastern hemlock [Tsuga canadensis (L.) Carr.]. P. 703-711 in Silvics of forest trees of the United States. Agriculture Handbook 271. USDA, Washington, D. C. FRANK, S.A. 1992. Models of plant-pathogen coevolution. Trends Genet. 8: 213-219. FRANKENHUYZEN, K. VAN, R. C. REARDON, AND N. R DUBOIS. 2000. Forest defoliators. P. 527-556 in Field manual of techniques in invertebrate pathology. Application and

170 evaluation of pathogens for control of insects and other invertebrate pests. Lacey L.A and H.K. Kaya (eds.). Kluwer Academic Pub. The Netherlands. FRASEN, J.J. 1987. Aschersonia aleyrodis as a microbial control agent of greenhouse whitefly. Ph. D. Thesis, Agricultural Univ. of Wageningen, The Netherlands. 167 p. FRASEN, J.J., AND J.C. VAN LENTEREN. 1993. Host selection and survival of the parasitoid Encarsia formosa on greenhouse whitefly, Trialeurodes vaporariorum, in the presence of hosts infected with the fungus Aschersonia aleyrodis. Entomol. Exp. Appl. 69: 239-249. FRASEN, J.J., AND J.C. VAN LENTEREN.1994. Survival of the parasitoid Encarsia formosa after treatment of parasitized greenhouse whitefly larvae with fungal spores of Aschersonia aleyrodis. Entomol. Exp. Appl. 71: 235-243. FRANSEN, J.J. 1995. Survival of spores of the entomopathogenic fungus Aschersonia aleyrodis (Deuteromycotina: Coelomycetes) on leaf surfaces. J. of Invertebrate Pathology 65: 73-75. FRASER J.A, S.S.GILES, E.C. WENINK, S.G. GEUNES-BOYER, J.R. WRIGHT, S. DIEZMANN, A. ALLEN, J.E. STAJICH, F.S. DIETRICH, J.R. PERFECT, AND J. HEITMAN. 2005. Same- sex mating and the origin of the Vancouver Island Cryptococcus gattii outbreak. Nature 437: 1360–1364. FREEMAN, S., AND R.J. RODRIGUEZ. 1992. A rapid, reliable bioassay for pathogenicity of Colletotrichum magna on cucurbits and its use in screening for nonpathogenic mutants. Plant Disease 76: 275-296. FREEMAN, S., AND R.J. RODRIGUEZ. 1995. Differentiation of Colletotrichum species responsible for anthracnose of strawberry by arbitrarily primed PCR. Mycol. Res. 99: 501-504. FREEMAN, S., T. KATAN, AND E. SHABI. 1996. Characterization of Colletotrichum gloeosporioides isolates from avocado and almond fruits with molecular and pathogenicity tests. Applied and Environmental Microbiology 62 (3): 1014-1020. FREEMAN, S., AND T. KATAN. 1997. Identification of Colletotrichum species responsible for anthracnose and root necrosis of strawberry in Israel. Phytopathology 87: 516- 521. FREEMAN, S., T. KATAN, AND E. SHABI. 1998. Host specificity and genetic diversity of Colletotrichum species on various crops. Workshop on host specificity, pathology and host – pathogen interaction of Colletotrichum. Ramat Rachel, Jerusalem, Israel. Phytoparasitica 26 (4): 5 (Abstr.). FREEMAN, S., D. MINZ, E. JURKEVITCH, M. MAUMON, AND E. SHABI. 2000. Molecular analyses of Colletotrichum species from almond and other fruits. Phytopathology 90: 608-614. FREEMAN, S., D. MINIZ, M. MAYMON, AND A. ZVEIBIL. 2001a. Genetic diversity within Colletotrichum acutatum sensu Simmonds. Phytopathology 91: 586-592. FREEMAN, S., S. HOROWITZ, AND A. SHARON. 2001b. Pathogenic and nonpathogenic lifestyles in Colletotrichum acutatum from strawberry and other plants. Phytopathology 91: 986-992. FUXA, J.R. 1987. Ecological considerations for the use of entomopathogens in IPM. Ann. Rev. Entomol. 32: 225-251.

171 FUXA, J.R., AND Y. TANADA. 1987. Epizootiology of insect diseases. Wiley Interscience Publ., NY. 555 p. GARRETT, W.T., AND G.S. LANGFORD. 1969a. Seasonal life cycle of Fiorinia externa in Maryland. J. Econ. Entom. 62 (5): 1221-1222. GARRETT, W.T. AND G.S. LANGFORD. 1969b. Control of Fiorinia externa, on hemlock in Maryland. J. Econ. Entom. 62 (6): 1449-1450. GILL, R.J. 1997. The scale insects of California Part 3. The Armored Scales (Homoptera: Diaspididae). California Dept. of Food and Agriculture, Sacramento, CA. 307 p. GOULI, V.V., S.Y. GOULI, S.D. COSTA, AND M.V. SHTERNSHIS. 2005. Comparison of wash, leaf imprint and adhesive tape methods for estimation of fungal spore deposition on leaves. Mycol. and Phytopath. 39 (3): 99-103. GLASS, N.L., AND G.A KULDAU. 1992. Mating type and vegetative incompatibility in filamentous ascomycetes. Ann. Rev. Phytopathol. 30: 201-224. GODMAN, R.M., AND K. LANCASTER. 1990. Tsuga canadensis (L.) Carr. In: Silvics of North America. Vol. 1. Conifers. R. M. Burns and B. H. Honkala (eds.). GOLD, S.E., M.D. GARCÍA-PEDRAJAS, AND A.D. MARTÍNEZ-ESPINOZA. 2001. New (and used) approaches to the study of fungal pathogenicity. Ann. Rev. Phytopathol. 39: 337-365. GONZÁLEZ, E., AND T.B. SUTTON. 2004. Population diversity within isolates of Colletotrichum spp. causing Glomerella leaf spot and bitter rot of apples in three orchards in North Carolina. Plant Disease 88: 1335-1340. GOSSARD, H.A. 1903. White Fly (Aleyrodes citri). Florida Agricultural Experiment Station Bulletin 67: 596-667. GOULI, S., B.L. PARKER, AND V. GOULI. 2004. Biological properties of a new entomopathogenic fungus Aschersonia marginata. P. 85 in Society for Invertebrate Pathology. 37th Annual Meeting. 7thInternational Conference on Bacillus thuringiensis. (Abstr.) GRAVENA, S., R.D.R LEAO-NETO, F.C. MORETTI, AND G. TOZATTI. 1988. Eficiência de insecticidas sobre Selenaspidus articulatus (Morgan) (Homoptera: Diaspididae) e efeito sobre inimigos naturais em pomar citrico. Cientifica (Jaboticabal) 16 (2): 209- 218. (In Portuguese) GRAVENA, S., P.T. YAMAMOTO, O.D. FERNANDES, AND I. BENETOLI. 1992. Efeito de Ethion e Aldicarb sobre Selenaspidus articulatus (Morgan), Parlatoria ziziphus (Lucas) (Homoptera: Diaspididae) e influencia sobre fungos patogėnicos. Anais da Sociedade Entomologica do Brasil 21 (2): 101-111. (In Portuguese) GRIFFIN, M. S., J. R SUTHEERELAND, AND J. J. DENNIS. 1987. Blight of conifer seedlings caused by Colletotrichum gloeosporioides. New Forest 1: 81-88. GRESSEL, J., D. MICHAELI, A. WARSHAWSKY, AND V. KAMPBEL 1998. Enhancing infectivity of a Colletotrichum mycoherbicide by chemical inhibiting callose biosynthesis in Workshop on host specificity, pathology and host – pathogen interaction of Colletotrichum. Ramat Rachel, Jerusalem, Israel. Phytoparasitica 26 (4): 11 (Abstr.)

172 GUARRO, J., T.E SVIDZINSKI, L. ZAROR, M. H. FORJAZ, J. GENÉ, AND O. FISCHMAN, 1998. Subcutaneous Hyalohyphomycosis caused by Colletotrichum gloeosporioides. J. of Clinical Microbiol. 36 (10): 3060-3065. GUERBER, J.C., AND J.C. CORRELL. 2001. Morphological description of Glomerella acutata, the teleomorph of Colletotrichum acutatum. Mycologia 93:216-229 GUERBER, J.C., B. LIU, J.C. CORRELL, AND P.R. JOHNSTON. 2003. Characterization of diversity in Colletotrichum acutatum sensu lato by sequence analysis of two gene introns, mtDNA and intron RFLPs, and mating compatibility. Mycologia 95 (5): 872- 895. GUOZHONG, L., P.F. CANNON, A. REID, AND C.M. SIMMONS. 2004. Diversity and molecular relationships of endophytic Colletotrichum isolates from the Iwokrama. Mycol. Res. 108: 53-63. GUNNELL, P.S., AND W.D. GUBLER. 1992. Taxonomy and morphology of Colletotrichum species pathogenic to strawberry. Mycologia 84 (2): 157-165. GUPTA, M., AND P. FILNER. 1991. Microsatellites amplify highly polymorphic DNA bands in SPAR of plant DNA. In: Proc. Int. Soc. Plant Mol. Biol., Tucson, AZ. (Abstr. 1705) GUZMAN, P., M.R. DONADO, AND G.E. GALVEZ. 1979. Perdidas economicas causadas por la antracnosis del frijol Phaeseolus vulgaris en Colombia. Turrialba 29: 65-67. HAJEK, A.E., J.S ELKINTON, AND J.J. WITCOSKY. 1996. Introduction and spread of the fungal pathogen Entomophaga maimaga (Zygomycetes: Enthomophthorales) along the leading edge of gypsy moth (Lepidoptera: Lymantriidae) spread. Biological Control 25 (5): 1236-1247. HALL, R.A., AND B. PAPIEROK. 1982. Fungi as biological control agents of arthropods of agricultural and medical importance. Parasitology 84: 205-240. HANAU, R.M, L.J. VAILLANCOURT, J. WANG, AND J.A. ROLLINS. 1998. Glomerella graminicola (Colletotrichum graminicola): evidence for a novel mechanisms for sex determination. Workshop on host specificity, pathology and host – pathogen interaction of Colletotrichum. Ramat Rachel, Jerusalem, Israel. Phytoparasitica 26 (4): 2 (Abstr.) HAWKSWORTH, D.L. 2001. The magnitude of fungal diversity: the 1.5 million species estimate revisited. Mycol. Res. 105 (12): 1422-1432. HELLE, W. 1990 a. Armored scale insects. Their biology, natural enemies and control. [Series title: World crop pests, Vol. 4A: The Armored scale insects]. Rosen, D. (ed.). Amsterdam, The Netherlands: Elsevier. 384 p. HELLE, W. 1990 b. Armored Scale Insects. Their biology, natural enemies and control. [Series title: World crop pests, Vol. 4B: The natural enemies]. Rosen, D. (ed.). Amsterdam, The Netherlands: Elsevier. 690 p. HILLIS, D.M., AND M.T. DIXON. 1991. Ribosomal DNA: molecular evolution and phylogenetic inference. Q. Rev. Biol. 66: 411–453. HOAGLAND, R.E. 1996. Chemical interactions with bioherbicides to improve efficacy. Weed Technology 10: 651-674. HOCH, H.C. 2004. Contact sensing: a tactilely important phenomenon for phytopathogenic fungi. Inoculum 55 (4): 17. (Abstr.)

173 HOOVER, G.A. 2003. Elongate hemlock scale. Entomological Notes. The Pennsylvania State Univ. www.ento.psu.edu/extension/ factsheets/pdfs/elongateHemlockScale.pdf HOPKINS, J.C., W. LOCK, AND A. FUNK. 1985. Colletotrichum acutatum, a new pathogen on western hemlock seedlings in British Columbia. Can. Plant Dis. Surv. 65: 11-13. HOROWITZ, S., S. FREEMAN AND A. SHARON. 2002. Use of green fluorescent protein- trangenic strains to study pathogenic and non-pathogenic lifestyles in Colletotrichum acutatum. Phytopathology 92: 743-749. HOROWITZ, S., O. YARDEN, A. ZVEIBIL, AND S. FREEMAN. 2004. Development of a robust screening method for pathogenicity of Colletotrichum spp. on strawberry seedlings enabling forward genetic studies. Plant Disease 88: 845-851. HOWARD, C.M., J.L. MAAS, C.K. CHANDLER, AND E.E. ALBREGTS. 1992. Antrachnose of strawberry caused by the Colletotrichum complex in Florida. Plant Disease 76 (10): 976-981. HOWARD, R.J., M.A. FERRARI, D.H. ROACH, AND N. P. MONEY. 1991. Penetration of hard substrates by a fungus employing enormous turgor pressure. PNAS 88: 11281-11284. HOWARD, T., P. SENDAK AND C. CODRESCU, 2000. Eastern hemlock: A market perspective. P. 161-166 in Proceedings: Symposium on sustainable management of hemlock ecosystems in eastern North America. McManus, K.A., K.S. Shields, and Souto, D.R. (eds). USDA Forest Service, Northeastern Research Station, Durham, NH. HOWE, R.W. AND M. MOSSMAN. 1995. The significance of hemlock for breeding birds in the Western Great Lakes region. P. 125-140 in Hemlock ecology and management. Proceedings of a regional conference on ecology and management of eastern hemlock. G. Mroz and A.J. Martin (eds.). Iron Mountain, MI. HUMBER, R.A. 1984. Foundations for an evolutionary classification of the Entomophthorales (Zygomycetes). P. 167-183 in Fungus-Insect relationships, Q. Wheeler, and M. Blackwell (eds.). Columbia Univ. Press, NY. HUMBER, R.A. 1989. Synopsis of a revised classification for the entomophthorales (Zygomycotina). Mycotaxon 34 (2): 441- 460. HUMBER, R.A. 1992. Collection of entomopathogenic fungi: Catalog of strains 1992. USDA-ARS Publications 110: 1-177. HUMBER, R.A. 1997. Fungi: identification. P. 153-185 in Manual of techniques in insect pathology, L.A. Lacey (ed.). Academic Press, London. HUMBER, R.A. 1998. Entomopathogenic fungal identification. APS/ESA Joint Annual Meeting. 26 p. HUMBER, R.A. 2000. Fungal pathogens and parasites of insects. P. 199-227 in Applied Microbial Systematics. F.G. Priest and M. Goodfellow (eds.). Kluwer Acad. Publ., Dordrecht. HUTCHISON, K.A., S.E. PERFECT, R.J. O’CONNEL, AND J.R. GREEN. 2000. Immunomagnetic purification of Colletotrichum lindemunthianum appressoria. Appl. and Env. Microbiology 66 (8): 3464-3467.

174 HIRTE, W.F., I. GLATHE AND H. ADAM. 1989. Produktion und application eines mikrobiellen Präparates auf der Basis von Aschersonia-Sporen. Zentralbl. Mikrobiol. 144: 151-162. (In German). HOPKINS, J.C., W. LOCK, AND A. FUNK. 1985. Colletotrichum acutatum, a new pathogen on western hemlock seedlings in British Columbia. Can. Plant Dis. Surv. 65: 11-13 IBRAHIM, Y.B. AND T.M. KON. 1992. In vivo pathogenicity of Aschersonia placenta and its telemorph on the scales, Asterolecanium ungulate and Coccus viridis. Malays. Appl. Biol. 21 (2): 71-75. IGNOFFO, C.M., AND R.F. ANDERSON. 1979. Bioinsecticides. P. 1-28 in Microbial Technology. Microbial Processes. Vol. 1. H.J. Peppler and D. Perlman (eds.). Academic Press, London. INGLIS, G.D., M.S. GOETTEL, T.M. BUTT, AND H. STRASSER. 2001. Use of Hypomycetous fungi for managing insect pests. P. 23-71 in Fungi as biocontrol agents. Butt, T.M., C. Jackson and N. Magan (eds). CAB International. INTERNATIONAL BIOHERBICIDE GROUP 2000. IBG News 9 (2): 1-18. IVEY, M.L.L., C. NAVA-DIAZ, AND S. A. MILLER. 2004. Identification and management of Colletotrichum acutatum on immature bell peppers. Plant Disease 88 (11): 1198- 1204. JACKSON, M.A., D.A. SCHISLER, AND B. S SHASHA. 1996. Formulation of Colletotrichum truncatum microsclerotia for improved biocontrol of the weed hemp sesbania (Sesbania exaltata). Biological Control 7 (1): 107-113. JAQUES, R.P. 1983. The potential of pathogens for pest control. Agriculture, Ecosystems and Environment 10: 101-126. JEFFRIES, P. 1998. Exploitation of microbial antagonism for the biocontrol of disease caused by Colletotrichum in Workshop on host specificity, pathology and host – pathogen interaction of Colletotrichum. Ramat Rachel, Jerusalem, Israel. Phytoparasitica 26 (4): 12 (Abstr.) JEPPSON, L.R., AND G.E. CARMAN. 1960. Citrus insects and mites. Ann. Rev. Entom. 5: 353-378. nd JOHNSON, W.T., AND H.H. LYON. 1998. Insects that feed on trees and shrubs. 2 Edition. Cornell Univ. Press. 556 p. JOHNSTON, P.R., AND D. JONES. 1997. Relationship among Colletotrichum isolates from fruit-rots assessed using rDNA sequences. Mycologia 89 (3): 420-430. DE JONG, J.C., B.J. MCCORMACK, N. SMIRNOFF, AND N.J. TALBOT. 1997. Glycerol generates turgor in rice blast. Nature 389: 244-245. JONSSON, C.M., AND F.J. GENTHNER. 1997. Avaliação do potencial de patogenicidade e toxicidade do fungo entomopatógeno Colletotrichum gloeosporioides isolados de Orthezia em duas espécies de crustáceos. Bol. Pesqui. Embrapa-CNPMA 1: 1-27. KELLER, S., C. SCHWEIZER, E. KELLER, AND H. BRENNER. 1997. Control of white grub (Melolontha melolontha L.) by treating adults with the fungus Beauvaria brongniartii. Biocontrol Science and Technology 7: 105-116. KELLER, S., AND G. ZIMMERMAN. 1989. Mycopathogens of Soil Insects. P. 239-270 in Insect-Fungus interactions. Wilding, N., N.M. Collins, P.M. Hammond and J.F. Webber (eds.). Academic Press. London.

175 KHAN, A., AND J.B. SINCLAIR. 1992. Pathogenicity of sclerotia - and nonsclerotia - forming isolates of Colletotrichum truncatum on soybean plants and roots. Phytopathology 82: 314-319. KHAN, A., AND T. HSIANG. 2003. The infection process of Colletotrichum graminicola and relative aggressiveness on four turfgrass species. Can. J. Microbiol. 49: 433-442. KILLGORE, E.M., L.S. SUGIYAMA, AND D.E. GARDNER. 1999. Evaluation of Colletotrichum gloeosporioides for biological control of Miconia calvescens in Hawaii. Plant Disease 83 (10): 964 (Abstr.) KIMPLE, A., AND W. SCHUSTER. 2002. Spatial patterns of Adelges tsugae damage and impacts on tree physiology and water use in the black rock forest, southern New York. P. 344-350 in Proceedings of the hemlock woolly adelgid in the eastern United States symposium. McManus, K.A., Shields, K. S. and Souto, D. R (eds.). USDA Forest Services, NJ. KISTLER, H.C. 1991. Genetic manipulation of plant pathogenic fungi. P. 152-170 in Microbial Control of Weeds. D.O. TeBeest (ed.). Routledge, Chapman & Hall, Inc. New York. 280 p. KOLATTUKUDY, P.E., Z. LIU, Y-K KIM, AND D. LI. 1998. Early molecular communication between Colletotrichum gloeosporioides and its hosts in Workshop on host specificity, pathology and host – pathogen interaction of Colletotrichum. Ramat Rachel, Jerusalem, Israel. Phytoparasitica 26 (4): 3 (Abstr.) KNAUER, K., J. LINNANE, K. SHIELDS, AND R. BRIDGES. 2002. An initiative for management of hemlock woolly adelgid. P. 9-11 in Proceedings of the hemlock woolly adelgid in the eastern United States symposium. McManus, K.A., Shields, K.S. and D.R. Souto (eds.). USDA Forest Services, NJ. KNIPLING, E.F. 1979. The basis principles of insect population supression and management. USDA. Agriculture handbook n. 512. Washigton. 659 p. KOSZTARAB, M. 1963. The armored scale insects of Ohio (Homoptera: Coccoidea: Diaspididae). Bulletin of the Ohio biological survey. Vol. II (2). 120 p. KOSZTARAB, M., AND F. KOZÁR. 1988. Scale insects of central Europe. Budapest: Akadémiai Kiadó [Joint Edition; also publ. in Dordecht, The Netherlands] 41: 456 p. KOSZTARAB, M., AND M.P. KOSZTARAB. 1988. A selected bibliography of the Coccoidea (Homoptera). Third supplement (1970-1985). Virginia polytechnic institute and state Univ. Agricultural experiment station bulletin 88-1: 1-252. KOSZTARAB, M. 1996. Scale insects of northeastern North America. Virginia Museum of Natural History special publication number 3. 645 p. KOVAL, E.Z. 1974. Identification of entomophilous fungi in the Soviet Union. Ukrainian Academy of Sciences. Kiev. 260 p. (In Russian) KRASNOFF, S.B., AND D.M. GIBSON. 1996. New destruxins from the entomopathogenic fungus Aschersonia sp. Journal of Natural Products 59: 485-489. KUBO, Y. 1998. Regulation of melanin biosynthesis genes during apressorium formation of Colletotrichum lagenarium. Workshop on host specificity, pathology and host – pathogen interaction of Colletotrichum. Ramat Rachel, Jerusalem, Israel. Phytoparasitica 26: 4 (Abstr.)

176 KULSHRESTHA, D.D., S.B MATHUR, AND P. NEERGAARD. 1976. Identification of seed- borne species of Colletotrichum. Contributions from the Danish government institute of seed pathology for developing countries. Friesia 40: 116-125. KUO, K-C. 1999. Germination and apressorium formation in Colletotrichum gloeosporioides. Proc. Natl. Sci. Counc. 23 (3): 126-132. KUWANA, A.M., S. ONUKI, AND S. HORI. 1904. The San José Scale in Japan. Imperial agriculture experiment station in Japan. Nishigahara, Tokyo. Tokyo Printing Co., Ltd. 33 p. LACEY, L.A., J.J. FRANSEN, AND R. CARRUTHERS. 1996. Global distribution of naturally occurring fungi of Bemisia, their biologies and use as biological control agents. P. 401-433 in: Bemisia 1995: Taxonomy, biology, damage, control and management. Gerling, D. and R. T. Mayer (eds.). Intercept Limited, Andover, UK. LAMBDIN P., C. LYNCH, J. GRANT, R. REARDON, B. ONKEN, AND R. RHEA. 2005. Elongate hemlock scale and its natural enemies in the southern Appalachians. P. 145-154 in 3rd symposium on hemlock woolly adelgid in the eastern United States, Onken, B., and R. Reardon (eds.). USDA Forest Service. LARDNER, R., K.M. PLUMMER, M.N. PEARSON, P.R JOHNSTON, AND M.D. TEMPLETON. 1997. Classical and molecular genetic analyses of Colletotrichum spp. 19th Fungal genetics conference. Von den Hondel, C. and G. Turgeon (eds.). Asilomar, California. (Abstr.) LARDNER, R., P.R. JOHNSTON, K.M. PLUMMER, AND M. N. PEARSON. 1999. Morphological and molecular analysis of Colletotrichum acutatum sensu lato. Mycol. Res. 103: 275-285. LATUNDE-DADA, A.O. 2001. Colletotrichum: tales of forcible entry, stealth, transient confinement and breakout. Molecular Plant Pathology 2 (4): 187-198. LEES, A.K., AND A.J. HILTON. 2003. Black dot (Colletotrichum coccodes): an increasingly important disease of potato. Plant Pathology 52: 3-12. LEGARD, D.E. 2000. Colletotrichum diseases of strawberry in Florida. P. 292-299 in Colletotrichum – Host specificity, pathology and host-pathogen interaction. Prusky, D., S. Freeman and M. B. Dickman (eds.). APS Press. St. Paul, MN. LENNÉ, J.M. 1992. Colletotrichum diseases of legumes. P. 135-166 in Colletotrichum – Biology, pathology and control. Bailey, J. A. and M. J. Jeger (eds.). CAB International. Wallingford, UK. LEWIS, L.C., D.J. BRUCK, R.D. GUNNARSON, AND K.G. BIDNE. 1996. Aptness of insecticides (Bacillus thurigiensis and carbofuran) with endophyc Beauvaria bassiana in suppressing larval populations of European corn borer. Agricultural Ecosystems and Environment 57: 27-34. LIM, T.K., Y.B. IBRAHIM, M.K. TANG, AND R. LIEW. 1991. Occurrence of Aschersonia placenta and Hypocrella raciborskii on Asterolecanium ungulate in Durian (Durio zibethinus). Biocontrol Science and Techonology 1: 137-144. LISANSKY, S.G., AND R. A. HALL. 1983. Fungal control of insects. P. 326-345 in The Filamentous Fungi, Vol. 4. J. E. Smith, D.R. Berry and B. Kristiansen (eds.). Fungal Technology. Edward Arnold, London.

177 LIU , B., AND J.C. CORRELL. 2000. Characterizaton of Colletotrichum species by analysis of a 1-kb intron of the glutamine synthetase gene. Phytopathology 90: S8. (Abstr.) LIU, B., J.C. GUERBER, AND J.C. CORRELL. 2001. Genetic diversity within and between subgroups of Colletotrichum acutatum based on RFLP and sequence analysis of a 1- kb intron of glutamine synthetase and a 200-bp intron of GPDH. Phytopathology 91: S55. (Abstr.) LIU, B. 2002. Molecular characterization and inter and intra species phylogenetic relationships in the fungal plant pathogen Colletotrichum. [Doctoral dissertation]. Univ. Arkansas. Fayetteville, Arkansas. 238 p. LOCK, W., J.R. SUTHERLAND, J.J. DENNIS, AND J.C. HOPKINS. 1984. Colletotrichum gloeosporioides shoot blight of greenhouse-grown western hemlock seedlings in British Columbia. Plant Disease 68: 628. LORD, J.C. 2005. From Metchnikoff to Monsanto and beyond: The path of microbial control. J. Invertebr. Pathol. 89: 19-29. LUBBE, C.M., S. DENMAN, S. CANNON, J.Z GROENEWALD, S.C. LAMPRECHT, AND P.W. CROUS, 2004. Characterization of Colletotrichum species associated with diseases of Proteaceae. Mycologia 96 (6): 1268-1279. MADDISON W.P., AND J.R. MADDISON. 1992. MacClade: Analysis of phylogeny and character evolution. Sinauer, Sunderland, MA. MAINS, E.B. 1959a. Species of Aschersonia. Lloydia. Vol. 22 (3): 215-221. MAINS, E.B. 1959b. North American species of Aschersonia parasitic on Aleyrodiidae. Journal of Invertebrate Pathology 1: 43-47. MAKOWSKI, R.M.D., AND K. MORTENSEN. 1992. The first mycoherbicide in Canada: C. gloeosporioides f. sp. malvae for round-leaved mallow control. P. 298-300 in Proc. of the 1st international weed control congress, Richardson, R. G. (ed.). Melbourne, Australia: Monarch University. MAKOWSKI, R.M.D., AND K. MORTENSEN. 1998. Latent infections and penetration of the bioherbicide agent Colletotrichum gloeospoprioides f. sp. malvae in non-target field crops under controlled environmental conditions. Mycol. Res. 102: 1545-1552. MALINOSKI, M.K., AND J.A. DAVIDSON. 2002. Scale insects on hemlock. Maryland cooperative extension entomology Leaflet # HG81. MANIRE, C.A., H.L. RHINEHART, D.A. SUTTON, E. H. THOMPSON, M.G. RINALDI, J.D. BUCK, AND E. JACOBSON. 2002. Disseminated mycotic infection caused by Colletotrichum acutatum in a Kemp’s ridley sea turtle (Lepidochelys kempi). J. of Clinical Microbiol. 40 (11): 4273-4280. MARRONE, P.G., AND S.C. MACINTOSH. 1993. Resistance to Bacillus thuringiensis and Resistance Management. P. 221-235 in Bacillus thuringiensis, An Environmental Biopesticide: Theory and Practice, Entwistle, P.F., J.S Cory, M.J. Bailey and S. Higgs (eds.). John Wiley & Sons, Chichester, UK. MARTIN, S., AND W. SKOROPAD. 1978. Nature of adhesive material of Colletotrichum graminicola appressoria. Trans. Br. Mycol. Soc. 10: 221-223. MARTÍN, M.P., AND F. GARCÍA-FIGUERES. 1999. Colletotrichum acutatum and C. gloeosporioides cause anthracnose in olives. Eur. J. Plant Pathol. 105: 733-741.

178 MARTINEZ-CULEBRAS, P.V., A. QUEROL, M.B. SUAREZ-FERNANDEZ, M.D. GARCIA- LOPEZ, AND E. BARRIO. 2003. Phylogenetic relationship among Colletotrichum pathogens of strawberry and design of PCR primers for their identification. J. Phytopathology 151: 135-143. MASEL, A.M., C. HE, A.M. POPLAWSKI, J.A.G. IRWIN, AND J.M. MANNERS. 1996 Molecular evidence for chromosome transfer between biotypes of Colletotrichum Mol. Plant-Microbe Interact. 9: 339–348. MAYER, M., R. CHIANESE, T. SCUDDER, J. WHITE, K. VONGPASEUTH, AND R. WARD. 2002. Thirteen years of monitoring the hemlock woolly adelgid in New Jersey forests. P. 50-60 in Proceedings of the hemlock woolly adelgid in the eastern United States Symposium, McManus, K.A., Shields, K. S. and Souto, D. R (eds.). New Jersey Agricultural Experiment Station, Rutgers University, New Brunswick, NJ. MCCLURE, M.S. 1977a. Resurgence of the scale, Fiorinia externa (Homoptera: Diaspididae) on hemlock following insecticide application. Environ. Entomol. 6: 480- 484. MCCLURE, M.S. 1977b. Dispersal of the scale Fiorinia externa (Homoptera: Diaspididae) and effects of edaphic factors on its establishment on hemlock. Environ. Entomol. 6: 539-544. MCCLURE, M.S. 1977c. Parasitism of the scale insect, Fiorinia externa (Homoptera: Diaspididae), by Aspidiotiphagus citrinus (Hymenoptera: Eulophidae) in a hemlock forest: Density dependence. Environ. Entomol. 6: 551-555. MCCLURE, M.S. 1977d. Ecology and control of Fiorinia externa Ferris (Homopetra: Diasppididae) on eastern hemlock. NY Ent. Soc. 85 (4): 187-188. MCCLURE, M.S., AND M.B FERGIONE. 1977. Fiorinia externa and Tsugaspidiotus tsugae (Homoptera: Diaspididae): Distribution, abundance and new hosts of two destructive scale insects of eastern hemlock in Connecticut. Environ. Entomol. 6: 807-811. MCCLURE, M.S. 1978a. Self regulation in scale insect population on hemlock. New York Entomological Society, Vol. 86 (4): 309. MCCLURE, M.S. 1978b. Seasonal development of Fiorinia externa, Tsugaspidiotus tsugae (Homoptera: Diaspididae), and their parasite, Aspidiotiphagus citrinus (Hymenoptera: Aphelinidae): Importance of parasite-host synchronism to the population dynamics of two scale pests of hemlocks. Environ. Entomol. 7: 863-870. MCCLURE, M.S. 1980a. Competition between exotic species: scale insects on hemlock. Ecology 61: 1391-1401. MCCLURE, M.S. 1980b. Foliar nitrogen: A basis for host suitability for elongate hemlock scale, Fiorinia externa (Homoptera: Diaspididae). Ecology 61: 72-79. MCCLURE, M.S. 1986. Population dynamics of Japanese hemlock scales: A comparison of endemic and exotic communities. Ecology 67 (5): 1411-1421. MCCLURE, M.S. 1988. The armored scales of hemlock. P. 46-65 in Dynamics of forest insect populations. A.A. Berryman (eds.). Plenum Press, New York. MCCLURE, M.S. 1997. Biological control in native and introduced species: Lessons learned from the sap-feeding guilds on hemlock and pine. P. 31-52 in Ecological Interactions and Biological Control. Andow D.A., D.W. Ragsdale and R. F. Nyvall (eds.). Westview Press, Boulder, Colorado.

179 MCCLURE, M.S. 2002a. The elongate hemlock scale, Fiorinia externa Ferris (Homoptera: Diaspididae): A new look at an old nemesis. P. 248-253 in Proceedings of the Hemlock woolly adelgid in the eastern United States symposium, Onken, B., R. Reardon and J. Lashomb (eds.). New Jersey Agricultural Experiment Station, Rutgers University, New Brunswick, NJ. MCCLURE, M.S. 2002b. Elongate hemlock scale. United States department of agriculture forest service pest alert. www.fs.fed.us/na/morgantown/fhp/palerts/ehs.pdf (last accessed in 5/6/06) MCCLURE, M.S., AND C.A.S.-J. CHEAH. 1999. Reshaping the ecology of invading populations of hemlock woolly adelgid, Adelges tsugae (Homoptera: Adelgidae), in eastern North America. Biological Invasions 1: 247-254. MCCOY, C.W. 1985. Citrus: current status of biological control in Florida. P. 481- 499 in Biological cControl in agricultural IPM Systems. Academic Press, London, NY. MCCOY, C.W., R.A. SAMSON, AND D.G. BOUCIAS.1988. Entomogenous fungi. P. 151- 236 in Handbook of natural pesticides, Vol. V. Microbial pesticides. Part A. Entomogenous protozoa and fungi, Ignoffo, C.M. and Mandava, N.B. (eds.), CRC Press, Boca Raton, FL. MCGUIRE D.A., AND R.T.T. FORMAN. 1983. Herb cover effects on tree seedling patterns in a mature hemlock-hardwood forest. Ecology 64: 1367-1380. MCRAE, C.F., AND B.A. AULD. 1988. The influence of environmental factors on anthracnose of Xanthium spinosum. Phytopathology 78: 1182-1186. MCWILLIAMS, W.H., AND T. SCHMIDT. 2000. Composition, structure, and sustainability of hemlock ecosystems in eastern North America. P. 5-10 in McManus, K.A., Shields, K. S. and Souto, D. R (eds.). USDA Forest Services, NJ. MEEKES, E.T.M., S. VAN VOORST, N.N. JOOSTEN, AND J.J. FRANSEN. 2000. Persistence of the fungal whitefly pathogen, Aschersonia aleyrodis, on three different plant species. Mycol. Res. 104 (10): 1234-1240. MEEKES, E.T.M., J.J. FRANSEN, AND J.C. VAN LENTEREN. 2002. Pathogenicity of Aschersonia spp. against whiteflies Bemisia argentifolii and Trialeurodes vaporariorum. Journal of Invertebrate Pathology 81: 1-11. MELOTTO, M., R.S. BALARDIN, AND J.D. KELLY. 2000. Host-Pathogen interaction and variability of Colletotrichum lindemunthianum. P. 346-361 in Colletotrichum – Host specificity, pathology and host-pathogen interaction, Prusky, D., S. Freeman and M. B. Dickman (eds.). APS Press. St. Paul, Minnesota. MEMMOTT, S.D., Y. HA, AND M.B. DICKMAN 2002. Proline reverses the abnormal phenotypes of Colletotrichum trifolii associated with expression of endogenous constitutively active Ras. Appl. and Env. Microbiology 68 (4): 1647-1651. MENDGEN, K., M. HAHN AND H. DEISING. 1996. Morphogenesis and mechanisms of penetration by plant pathogenic fungi. Ann. Rev. Phytopathol. 34: 367-386. MERTELY, J.C. AND D. E. LEGARD. 2004. Detection, isolation and pathogenicity of Colletotrichum spp. from strawberry petioles. Plant Disease 88: 407-412. METAMORPH IMAGING SYSTEM (2003). Meta imaging series MetaMorph basic commands: Version 6.1 for Microsoft Windows 2000/XP: User's Guide. Universal Imaging Corporation, Downingtown, PA 19355.

180 MILLER, J.H. 1940. The genus Myriangium in North America. Mycologia 32: 587-600. MILLS, P.R., S. SREENIVASAPRASAD, AND A.E. BROWN. 1992. Detection and differentiation of Colletotrichum gloeosporioides isolates using PCR. FEMS Microbiology Letters 98: 137-144. MILLS, P.R., S. SREENIVASAPRASAD, AND A.E. BROWN. 1994. Detection of the antracnose pathogen Colletotrichum. P. 183-189 in Modern assays for plant pathogenic fungi. Schots, S., F.M Dewey and R. P. Oliver (eds.). CAB International. Wallingford, UK. MINDELL, D.P. AND R. HONEYCUTT. 1990. Ribosomal RNA in vertebrates: Evolution and phylogenetic applications. Ann. Review of Ecology and Systematics 21: 541-566. MLADENOFF, D.J. 1995. The role of eastern hemlock across scales in the northern lake states. P. 29-42 in Hemlock ecology and management. Proceedings of a regional conference on ecology and management of eastern hemlock. Mroz, G. and A. J. Martin (eds.). Dept. of Forestry, University of Wisconsin, Madison. MONEY, N.P. 1995. Turgor pressure and the mechanics of fungal penetration. Can. J. Bot. 73: S96-S102. MONEY, N.P. 1999a. Fungus punches its way in. Nature 401: 332-333. MONEY, N.P. 1999b. To perforate a leaf of grass. Fungal Genet. Biol. 28: 146-147. MONTAGNE, C. 1848. Aschersonia Montag. Nov. Gen. Ann. Sci. Nat. III. 10: 121. MORETTO, K.C.K., N. GIMENES-FERNANDES AND J.M. SANTOS. 2001. Influence of Trichoderma spp. on Colletotrichum acutatum mycelial growth and morphology and on infection of ‘Tahiti’ lime detached flowers. Summa Phytopathologica 27: 357-364. (In Portuguese) MORIN, L., J-A.L. DERBY, AND E.G. KOKKO. 1996. Infection process of Colletotrichum gloeosporioides f. sp. malvae on Malvaceae weeds. Mycol. Res. 100 (2): 165-172. MORRIS, M.J. 1983. Evaluation of field trials with Colletotrichum gloeosporioides for the biological control of Hakea sericea. Phytophylactica. 15 (1): 13-16. MORRISON, H., AND A.V. RENK. 1957. A selected bibliography of the Coccoidea. Miscellaneous publications United States department of agriculture 734: 1-222. MORRISON, H., AND E.R. MORRISON. 1965. A selected bibliography of the Coccoidea. First Supplement (1956-1962). Miscellaneous publications United States department of agriculture 987: 1-44. MORRISON, K.D., E.G. REEKIE, AND K.I.N. JENSEN. 1998. Biocontrol of common St. Johnswort (Hypericum perforatum) with Chrysolina hyperici and a host-specific Colletotrichum gloeosporioides. Weed Technology 12: 426-435. MORIWAKI, J., T. TSUKIBOSHI, AND T. SATO. 2002. Grouping of Colletotrichum species in Japan based on rDNA sequences. J. Gen. Plant Pathol. 68: 307-320. MORTENSEN, K. 1988. The potential of an endemic fungus, Colletotrichum gloeosporioides, for biological control of round-leaved mallow (Malva pusilla) and Velvetleaf (Abutilon theophrasti). Weed Science 36: 473-478. MOTULSKY, HJ. 1999. Analyzing Data with GraphPad Prism. GraphPad Software Inc. San Diego, CA . 193 p. MUELLER, G.M., G.F. BILLS, AND M.S. FOSTER. 1995. Biodiversity of fungi. Inventory and monitoring methods. Elsevier Academic Press. 728 p.

181 MUNIZ, M. DE F. S., R. DE C. R. SANTOS AND G.V. DE S. BARBOSA. 1998. Pathogenicity of Colletotrichum gloeosporioides isolates on some tropical fruits. Summa Phytopathologica 24: 177-179. (In Portuguese) NACHET, K.L., AND M.S. ABREU. 2002. Morphological characterization and pathogenicity of isolates of Colletotrichum sp. from coffee. Ciênc. Agrotec., Lavras 26 (6): 1135- 1142. NAIR, J. AND J.B. CORBIN. 1981. Histopathology of Pinus radiata seedlings infected by Colletotrichum acutatum f. sp. pinea. Phytopathology 71: 777-783. NAIR, J., F.J. NEWHOOK, AND J.B. CORBIN. 1983. Survival of Colletotrichum acutatum f. sp. pinea in soil and pine debris. Trans. Br. Mycol. Soc. 81 (1): 53-63. NEGASI, A., B.L. PARKER, AND M. BROWNBRIDGE. 1998. Screening and bioassay of entomopathogenic fungi for the control of silverleaf whitefly, Bemisia argentifolii. Insect Sci. Applic. 18: 37-44. NICHOLSON, R.L. 1992. Colletotrichum graminicola and the anthracnose disease of maize and sorghum. P. 186-202 in Colletotrichum – Biology, pathology and control, Bailey, J.A. and M.J.Jeger (eds.). CAB International. Wallingford, UK. NIRENBERG, H.I., U. FEILER, AND G. HAGEDORN. 2002. Description of Colletotrichum lupini comb. nov. in modern terms. Mycologia 94 (2): 307-320. NIKOH, N., AND T. FUKATSU. 2000. Interkingdom host jumping underground: phylogenetic analysis of entomoparasitic fungi of the genus Cordyceps. Mol. Biol. Evol. 17: 629-638. NOF, E., B. RUBIN, AND A. DINOOR. 1998. Colletotrichum sp. - A potential, specific biocontrol agent for dodder (Cuscuta campestris). Workshop on host specificity, pathology and host – pathogen interaction of Colletotrichum. Ramat Rachel, Jerusalem, Israel. Phytoparasitica 26 (4): 11 (Abstr.) NORMAN, D.J., AND J.O. STRANDBERG. 1997. Survival of Colletotrichum acutatum in soil and plant debris of leatherleaf fern. Plant Disease 81: 1177-1180. O’DONNELL, K. 1992. Ribosomal DNA internal transcribed spacers are highly divergent in the phytopathogenic ascomycete Fusarium sambucinum (Gibberella pulicaris). Curr. Genet. 22: 213–220. O’DONNELL, K. 1993. Fusarium and its near relatives. P. 225–236 in The fungal holomorph: mitotic, meiotic, and pleomorphic speciation in fungal systematics, Reynolds, D.R., and J.W. Taylor. (eds.). CAB International, Wallingford, UK. OLIVER, R.P., AND S.V.S. IPCHO. 2004. Arabidopsis pathology breathes new life into the necrotrophs-vs.-biotrophs classification of fungal pathogens. Molecular Plant Pathology 5 (4): 347-352. ORWIG, D.A., AND D.R. FOSTER. 1998. Forest response to the introduced hemlock woolly adelgid in southern New England, USA. Journal of the Torrey Botanical Society 125 (1): 60-73. ORWIG, D.A. 2002. Stand dynamics associated with chronic hemlock woolly adelgid infestations in southern New England. P. 36-46 in Proceedings of the hemlock woolly adelgid in the eastern United States symposium, Onken, B., R. Reardon and J. Lashomb (eds.). New Jersey Agricultural Experiment Station, Rutgers University, New Brunswick, NJ.

182 ORWIG, D.A., D.R. FOSTER, AND D.L. MAUSEL. 2002. Landscape patterns of hemlock decline in New England due to the introduced hemlock woolly adelgid. Journal of Biogeography 29: 1475-1487. OSBURN, M.R. AND H. SPENCER. 1938. Effect of spray residues on scale insect populations. Journal of Economic Entomology 31 (6): 731-732. PANNACIONE, D.G, AND R.M. HANAU. 1990. Characterization of two divergent ß-tubulin genes from Colletotrichum graminicola. Gene 86: 163-170. PARKER, B.L., M. SKINNER, V. GOULI, S. GOULI, J. MARCELINO, J. CARLSON AND L. SCHARTZBERG. 2005. Management of elongate hemlock scale with entomopathogenic fungi. P. 161-168 in 3rd Symposium on hemlock woolly adelgid in the eastern United States, Onken B., and R. Reardon (eds.). USDA Forest Service. PARKIN, J. 1906. Fungi parasitic upon scale-insects (Coccidae and Aleurodidae). Ann. Roy. Bot. Gard. (Peradeniya) 3: 11-82. PARIKKA, P., AND A. LEMMETTY. 2004. Tracing latent infection of Colletotrichum acutatum on strawberry by PCR. European Journal of Plant Pathology 110: 393-398. PEREIRA, J.C.R., U.G. BATISTA, F.B. GUIMARÃES, AND E.S.G. MISUBUTI. 1998. Efeito de diferentes meios de cultura sobre a esporulação e o potencial de inóculo de Colletotrichum lindemuthianum. Summa Phytopathologica 24: 186-189. (In Portuguese) PERES N.A., L.W. TIMMER, J.E. ADASKAVEG, AND J.C. CORRELL. 2005. Lifestyles of Colletotrichum acutatum. Plant Dis 89: 784-796. PERFECT, E.S., AND J.R. GREEN. 2001. Infection structures of biotrophic and hemibiotrophic fungal plant pathogens. Mol. Plant Pathol. 2: 101-108. PERKINS, D.D. 1987. Mating type switching in filamentous ascomycetes. Genetics 115: 215-216. PERKINS, J.H. 1988. Naturally occurring pesticides and the pesticide crisis. 1945 to 1980. P. 151-236 in Handbook of natural pesticides. Vol. V. Microbial insecticides, Part A, Entomogenous, protozoas and fungi. C.M. Ignoffo and N.B. Mandava (eds.). CRC Press. PETCH, T. 1914. The genera Hypocrella and Aschersonia (a preliminary note). Ann. Roy. Bot. Gard. (Peradeniya) 5: 521-537. PETCH, T. 1921. Studies on entomogenous fungi. II. The genera Hypocrella and Aschersonia. Ann. Roy. Bot. Gard. (Peradeniya) 7: 167-278. PETCH, T. 1922. Studies on entomogenous fungi. Trans. Brit. Mycol. Soc. Cambridge 33 (2-3): 89-167. PETCH, T. 1925. Entomogenous fungi: additions and corrections. Transactions of the British Mycological Society. 10: 190-201. PETCH, T. 1927. Entomogeneous fungi – additions and corrections. Trans. Brit. Myc. Soc. pt. 3-4: 258-266. POINAR, G.O., AND G.M. THOMAS. 1978. Diagnostic manual for the identification of insect pathogens. Plenum Press, New York and London. 218 p. POINAR, G.O., AND G.M. THOMAS. 1982. An entomophtoralean fungus from Dominican amber. Mycologia 74: 332-334.

183 POINAR, G.O., AND G.M. THOMAS. 1984. A fossil entomogenous fungus from Dominican amber. Experientia 40: 578-579 PONOMARENKO, N.G., N.A. PRILEPSKAIA, M.J.A. MURVANIDZE, AND L.A. STOLYAROVA, 1975. Aschersonia for control of the citrus whitefly Dialeurodes citri. Zashch. Rast. 6: 44-45. (In Russian) PRUSKY, D., AND R.A. PLUMBEY. 1992. Quiescent infection of Colletotrichum in tropical and subtropical fruits. P. 289-307 in Colletotrichum – Biology, pathology and control, Bailey, J. A. and M. J. Jeger (eds.). CAB International. Wallingford, UK. PRUSKY, D., I. KOBILER, R. ARDI, D. BENO-MOALEM, AND N. YAKOBY. 1998. Mechanisms of defense of unripe avocado fruits against Colletotrichum gloeosporioides attack in Workshop on host specificity, pathology and host – pathogen interaction of Colletotrichum. Ramat Rachel, Jerusalem, Israel. Phytoparasitica 26 (4): 10 (Abstr.) PRUSKY, D., S. FREEMAN, AND M. B. DICKMAN. 2000. Colletotrichum – Host specificity, pathology and host-pathogen interaction, Prusky, D., S. Freeman, and M. B. Dickman (eds.). APS Press. St. Paul, Minnesota. 393 p. PRUSKY, D., J.L. MCEVOY, AND W. S. CONWAY. 2002. Local pH modulation by pathogens as a mechanism for increase virulence. P. 34 in 6th European Conference on Fungal Genetics. Pisa, Italy. (Abstr.) RAID, R.N., AND S.P. PENNYPACKER. 1987. Weeds as hosts for Colletotrichum coccodes. Plant Disease 71: 643-646. RAMAKERS, P.M.J., AND R.A. SAMSON. 1984. Aschersonia aleyrodis, a fungal pathogen of whitefly. Zeitschrift fur angewandte Entomologie 97: 1-8. RAND, F.V. 1914. Some diseases of pecan. J. Agric. Res. 1: 303-338. RAO, N.N.R., AND H.S. SOHI. 1980. Occurrence of Podonectria coccicola (Ellis And Everh.) Petch on citrus scale insects. Current Science 49 (9): 359-360. REDMAN, R.S., D.D. DUNIGAN, AND R.J. RODRIGUEZ. 2001. Fungal symbiosis from mutualism to parasitism: who controls the outcome, host or invader?. New Phytopatologist 151: 705-716. REITZ, S.R., AND J.T. TRUMBLE. 2002. Competitive displacement among insects and arachnids. Ann. Rev. Entomol. 47: 435-465. RITTERBAND, D.C., M. SHAH, AND J.A. SEEDOR. 1997. Colletotrichum graminicola: a new corneal pathogen. Cornea 16: 362-364. ROANE, K.M., G.J. GRIFFIN, AND J.R. ELKINS. 1986. Chestnut Blight, other Endothia diseases, and the genus Endothia. APS Press. St. Paul, EUA. ROBBS, C.F., L.A.N. SÁ, F. LUCCHINI, R. CESNIK, AND C.V.S. SADI. 1990. Control of scale Orthesia praelonga Douglas in Citrus orchards with entomopathogenic strains of Colletotrichum gloeosporioides Penz. P. 118 in Anais do 2º Simpósio de controle biológico. Brasília, Brazil. (In Portuguese) ROBBS, C.F., L.A.N. SÁ, F. LUCCHINI, R. CESNIK, AND C.V.S. SADI. 1991. Utilização de estirpes do fungo Colletotrichum gloeosporioides no controle da Orthezia praelonga. P. 245 in 13o Congresso brasileiro de entomologia. Recife, Brazil. (In Portuguese)

184 ROBBS, C.F., R. CESNIK, C.V.S., SADI, L.A.N SÁ, AND F. LUCCHINI. 1993. Controle biológico da Orthezia praelonga. P. 305 in 15º Congresso brasileiro de entomologia. Piracicaba, Brazil. (In Portuguese) ROBERTS, D.W., AND S.P. WRAIGHT. 1986. Current status on the use of insect pathogens as biocontrol agents in agriculture: Fungi. P. 510-513 in Fundamental and applied aspects of invertebrate pathology. Samson, R.A., Vlak, J.M. and Peters, D. (eds.). Published by the Foundation of the fourth international colloquium of invertebrate pathology. Wageningen, The Netherlands. ROBERTS, D.W. AND R.A. HUMBER. 1981. Entomogenous fungi. P. 201-236 in Biology of Conidial Fungi. T.C. Garry and B. Kendrick (eds.). ROBERTS, D.W. 1981. Toxins of entomopathogenic fungi. P. 201-236 in Microbial control of pests and plant diseases 1970-1980. H. D. Burges (ed.). Academic Press. New York. ROBERTS, D.W., AND R.A. HUMBER. 1984. Entomopathogenic fungi. P. 1-12 in Infection processes of fungi. Roberts, D. W. and J. R. Aist (eds.). Publs. Rockefeller Fdn. NY. ROCA, M., AND R.S. REDMAN. 2000. Colletotrichum as a model system for defining the genetic basis of fungal symbiotic lifestyles. P. 114-130 in Prusky D., S. Freeman, M.B. Dickman (eds.). Colletotrichum – Host specificity, pathology and host-pathogen interaction. St. Paul, Minnesota: APS Press. ROCA, M.G., J. MACHADO, M.G.G.C. VIEIRA, L.C. DAVIDE, AND M.L.M. ROCHA. 2004a. Compatibilidade sexual e vegetativa do complemento Glomerella-Colletotrichum associado a sementes de algodão. Fitopatol. Bras. 29 (1): 16-20. (In Portuguese) ROCA, M.G., L.C. DAVIDE, L.M.C. DAVIDE, M.C. MENDES-COSTA, R.F. SCEN, AND A.E. WHEALS. 2004b. Conidial anastomosis fusion between Colletotrichum species. Mycol. Res. 108 (11): 1320-1326. RODGERS, P.B. 1993. Potential of biopesticides in agriculture. Pesticide Science 39: 117- 129. RODRIGUEZ, R., R.S. REDMAN, AND J. RANSON. 1998. Defining the genetics of host specificity and symbiotic life styles in Colletotrichum species. Workshop on host specificity, pathology and host – pathogen interaction of Colletotrichum. Ramat Rachel, Jerusalem, Israel. Phytoparasitica 26 (4): 5 (Abstr.) RODRIGUEZ, R.J., AND R.S. REDMAN. 2000. Colletotrichum as a model system for defining the genetic basis of fungal symbiotic lifestyles. P. 114-130 in Colletotrichum – Host specificity, pathology and host-pathogen interaction. Prusky, D., S. Freeman and M. B. Dickman (eds.). APS Press. St. Paul, Minnesota. ROJAS, T. 2000. Aschersonia basicystis sobre insectos escama (Homoptera: Coccidae) en Venezuela. Rev. Iberoam. Micol. 17 (4): 135-137. ROLFS, P.H. 1897. A fungus disease of the San Jose scale. Sphaerostilbe coccophila, Tul. Florida Agricultural Experiment Station. Bulletin 41: 516-542. ROLFS, P.H., AND H.S. FAWCETT. 1908. Fungus diaseases of scale-insects and whiteflies. Florida Agricultural Experiment Station. Bulletin 94: 1-17. ROMBACH, M.C., AND A.T. GILLESPIE. 1988. Entomogenous Hyphomycetes for insect and mite control on greenhouse crops. Biocontrol News and Information. Vol. 9 (1): 7-18.

185 ROSAMIGLIA, A.C., AND I.S. MELO. 1996. Caracterização morfológica e bioquímica de Colletotrichum gloeosporioides, agente de controle biológico de Orthezia praelonga. P. 399 in Quinto simpósio de controle biológico. Foz de Iguaçu, Brazil. (In Portuguese). (Abstr.) ROSE, M. 1990. Rearing and mass rearing. P. 357-365 in Armored scale insects. Their biology, natural enemies and control. [Series title: World crop pests, Vol. 4A: The armored scale insects], Rosen, D. (ed.). Elsevier, Amsterdam. ROSSMAN, A.Y. 1994. A strategy for an all-taxa incentory of fungal diversity. P. 169-194 in Biodiversity and terrestrial ecosystems. C.-I. Peng and Chen C.H. (eds.). [Monograph Series No.14]. Institute of Botany, Academia Sinica, Taipei. ROYLE, D.D., AND R.G. LATHROP. 2002. Using Landsat imagery to quantify temporal and spatial patterns in hemlock decline. P. 67-72 in Proceedings of the hemlock woolly adelgid in the eastern United States symposium. Onken, B., R. Reardon and J. Lashomb (eds.). New Jersey Agricultural Experiment Station, Rutgers University, New Brunswick, NJ. RUSSELL, L.M., M. KOSZTARAB, AND M.P. KOSZTARAB. 1974. A selected bibliography of the Coccoidea. Second supplement (1963-1969). Miscellaneous publications United States department of agriculture 1281: 1-122. SASSCER, E.R. 1912. The genus Fiorinia in the United States. USDA Bur. Entomol. Tech. Ser. 16: 75-82. SAMSON, R.A. AND C.W. MCCOY. 1983. Aschersonia aleyrodis, a fungal pathogen of whitefly. I. Scanning electron microscopy of the development on the citrus whitefly. Z. ang. Ent. 96: 380-386. SAMSON, R.A., H.C. EVANS, AND J-P. LATGÉ. 1988. Atlas of entomopathogenic fungi. Springer-Verlag, Berlin. 187 p. SANDS, D.P.A., AND Van R.G. DRIESCHE. 2000. Evaluating host specificity of agents for biological control of arthropods: rationale, methodology and interpretation. P. 69– 83 in 10th International symposium on biological control of weeds, Van Driesche, R.G., T.A, Heard, A.S., McClay, and R. Reardon (eds.). Forest Service Bulletin FHTET- 99-1, Morgantown, WV. SANDERS, G.M., AND L. KORSTEN. 2003. A comparative morphological study of South African avocado and mango isolates of Colletotrichum gloeosporioides. Can. J. Bot. 81: 877-885. SANOGO, S., AND S.P. PENNYPACKER. 1997. Factors affecting sporogenic and myceliogenic germination of sclerotia of Colletotrichum coccodes. Plant. Disease 81: 333-336. SARMIENTO, M.M.V., D.A.R SIERRA, J. SANABRIA, AND A. LOPEZ-AVILA. 1995. Ensayo de diferentes dosis de Aschersonia aleyrodis Webber y parasitismo de Encarsia Formosa Gahan en ninfas de tercer e cuarto instar de la mosca blanca de los invernaderos. Revista Colombiana de Entomologia. Vol. 21 (3): 159-170. (In Spanish) SAS INC. 1990. SAS/STAT user's guide. 4th ed. Cary, NC. SAS Institute. 1069 p.

186 SCHMITTHENNER, A.F., AND J.W. HILTY. 1962. A modified dilution technique for obtaining single-spore isolates of fungi from contaminated material. Phytopathology 52: 582–583. SEAVER, F.J. 1920. Notes on North American Hypocreales – IV. Aschersonia and Hypocrella. Mycologia 12: 93-98. SHARON, A. 1998. The Potential of weed-pathogenic Colletotrichum species as mycoherbicides in Workshop on host specificity, pathology and host – pathogen interaction of Colletotrichum. Ramat Rachel, Jerusalem, Israel. Phytoparasitica 26 (4): 10 (Abstr.). SHARON A., K. YAMAGUCHI, S. CHRISTIANSEN, B.A. HORWITZ, O.C YODER, AND B.G. TURGEON. 1996. An asexual fungus has the potential for sexual development. Mol. Gen. Genet. 251: 60-68. SHAH, P.A. AND J.K. PELL. 2003. Entomoparthogenic fungi as biological control agents. Appl. Microbiol. Biotechnol. 61: 413-423. SHERRIFF, C., M.J. WHELAN, G.M. ARNOLD, J-F LAFAY, Y. BRYGOO, AND J.A. BAILEY 1994. Ribosomal DNA sequence analysis reveals new species groupings in the genus Colletotrichum. Experimental Mycology 18: 121-138. SHIRYAEVA, N.V. 2000. A dangerous pest on the Black Sea coast of the Caucasus. Zashchita I Karantin Rastenii 12: 33. SHOEMAKER, R.A. 1981. Changes in taxonomy and nomenclature of important genera of plant pathogens. Ann. Rev. Phytopathol. 19: 297-307 SHUTOVA, N.N., A.I. ANTADZE, M.N. MURVANIDZE, AND T.V. TIMOFEEVA. 1985. Biological control of pests of citrus crops. Zashchita Rastenii 2: 37. SILMAN, R.W., R.J. BOTHAST, AND D.A. SCHISLER. 1993. Production of Colletotrichum truncatum for use as a mycoherbicide: effects of culture, drying and storage on recovery and efficacy. Biotech. Adv. 11: 561-575. SIMMONDS, J.H. 1965. A study of the species of Colletotrichum causing ripe fruit rots in Queensland. Queensland Journal of Agricultural and Animal Science 22: 437-459. SINCLAIR, J.B. 1991. Latent infection of soybean plants and seeds by fungi. Plant Disease 75: 220-224. SLADE, S.J., T.R. SWINBURNE, AND S.A. ARCHER. 1986. The role of a bacterial siderophore and of iron in the germination and apressorium formation by conidia of Colletotrichum acutatum. J. of General Microbiology 132: 21-26. SMITH, A.H. 1969. Edwin Butterworth Mains, 1890-1968. Mycologia 61 (3):449-451. SMITH, B.J., AND L.L. BLACK. 1986. First report of Colletotrichum acutatum on strawberry in the United States. Plant Disease 70 (11): 1074. SMITH, B.J. 1990. Morphological, cultural, and pathogenicity variation among Colletotrichum species isolated from strawberry. Plant Disease 74 (1): 69-76. SMITH, D., D. PAPECEK, AND N. SMITH. 1995. Biological control of citrus snow scale, Unaspis Citri (Comstock) (Homoptera: Diaspididae) in South-East Queensland, Australia in Proceedings of the VII international symposium of scale insect studies. The Volcani Center, Agricultural Research Organization, Bet Dagan, Israel. Israel Journal of Entomology. Vol. 24: 253-260.

187 SMITH, J.E., AND T.A.S.A. AVELING. 1997. Colletotrichum demantium: Causal agent of a new cowpea stems disease in South Africa. Plant Disease 81 (7): 832. SMITH, J.E., L.K. KORSTEN, AND T.A.S.A. AVELING. 1999. Infection process of Colletotrichum demantium on cowpea stems. Mycol. Res. 103 (2): 230-234. SMITH, R. J. 1991a. Integration of biological control agents with chemical pesticides. P. 189-208 in Microbial control of weeds. D.O. TeBeest (ed.). Routledge, Chapman and Hall, Inc. New York. SMITH, V.L. 1991b. Canker of grafted Japanese maple caused by Colletotrichum acutatum. Phytopathology 81: 1177. (Abstr.) SMITH, V.L. 1993. Canker of Japanese maple caused by Colletotrichum acutatum. Plant Disease 77: 197-198. SMITH, K.T., AND P.A. LEWIS. 2005. Potential concerns for tree wound response from stem injections. P. 173-178 in 3rd Symposium on hemlock woolly adelgid in the eastern United States. Onken B., and R. Reardon (eds.). USDA Forest Service. SNYDER C.D., J.A. YOUNG, D.P. LEMARIE, AND D.R. SMITH. 2002. Influence of eastern hemlock (Tsuga canadensis) forests on aquatic invertebrate assemblages in headwater streams. Can. J. Fish. Aquat. Sci. 59(2): 262-275. SOUTO, D.R., AND K.S. SHIELDS. 2000. Overview of hemlock health. P. 76-80 in Proceedings symposium on sustainable management of hemlock ecosystems in eastern North America. McManus, K.A., Shields, K. S. and Souto, D. R (eds.). USDA Forest Service, Northeastern Research Station, Durham, NH. SOLOVEI, E.F., AND P.D. KOLTSOV. 1976. The action of entomogenous fungi of the genus Aschersonia on the whitefly. Mikol. Fitopatol. 10 (5): 425-429. (In Russian) SOUTH, F.W. 1910. The control of scale insects in the British West Indies by means of fungoid parasites. West Indian Bulletin 11: 1-30. SPASSOVA, P. 1980. Investigations on the entomopathogenic fungus Aschersonia sp. Gradinarska I Lozarska Nauka 17 (3/4): 77-83. (In Russian) SPASSOVA, P., E. HRISTOVA, AND E.S. ELENKOV. 1980. Pathogenicity of various species of fungi belonging to the genus Aschersonia to the larvae of glasshouse whitefly (Trialeurodes vaporiorum Westwood) on tomatoes and cucumbers. Hortic. Vitic. Sci. XVII (5): 70-76. (In Russian) SPATAFORA, J.W., G-H. SUNG, J-M SUNG, N.L. HYWEL-JONES, AND F. WHITE. 2007. Phylogenetic evidence for an animal pathogen origin of ergot and the grass endophytes. Mol. Ecol. 16: 1701-1711. SPSS Inc. 2005. SPSS Base 14.0 for Windows User's Guide. Chicago, IL. SPSS Inc. 783 p. SREENIVASAPRASAD, S., A. BROWN, AND P. R. MILLS. 1993. Coffee berry disease pathogen in Africa: genetic structure and relationship to the group species Colletotrichum gloeosporioides. Mycol. Res. 97 (8): 995-1000. SREENIVASAPRASAD, S., P.R. MILLS, B.M. MEEHAN, AND A.E BROWN. 1996. Phylogeny and systematics of 18 Colletotrichum species based on ribosomal DNA spacer sequences. Genome 39: 499-512.

188 SREENIVASAPRASAD, S., AND P. TALHINHAS. 2005. Pathogen profile. Genotypic and phenotypic diversity in Colletotrichum acutatum, a cosmopolitan pathogen causing anthracnose on a wide range of hosts. Molecular Plant Pathology 6 (4): 361-378. STEINHAUS, E.A. 1949. Principles of insect pathology. McGraw-Hill Book Company, Inc. 710 p. STIMMEL, J.F. 1980. Seasonal history and occurrence of Fiorinia externa Ferris in Pennsylvania (Homoptera: Diaspididae). Proc. Entomol. Soc. Wash. 82 (4): 700-706. STURM, J. 1831. ‘Colletotrichum’ Corda. Deutschlands flora in abbildungen nach der natur mit beschreibungen. III. Abtheilung. Die pilze Deutschlands: 41. (In Latin and German). SHIRYAEVA, N.V. 2000. A dangerous pest on the black sea cost of the Caucasus. Zashchita i Karantin Rasteni 12: 33. (Abstr.) STANISLAW, B. 1999. Monograph fungi. Polish Academy of Sciences. 353 p. (In Russian) STEPHENSON S.A., J.R. GREEN, J.M. MANNERS, AND D.J. MACLEAN. 1997. Cloning and characterization of glutamine synthetase from Colletotrichum gloeosporioides and demonstration of elevated expression during pathogenesis on Stylosanthes guianensis. Curr. Genet. 31: 447-54. STOETZEL, M.B. 1976. Scale-cover formation in the Diaspididae (Homoptera: Coccoidea). Proc. Ent. Soc. Wash. 78: 323-332. SUTTON, B.C. 1992. The genus Glomerella and its anamorph Colletotrichum. P. 1-26 in Colletotrichum – Biology, Pathology and Control. Bailey, J. A. and M. J. Jeger (eds.). CAB International. Wallingford, UK. SUTTON, B.C. 1968. The appressoria of Colletotrichum graminicola and C. falcatum. Can. J. Bot. 46: 873-876. SWOFFORD, D.L. 2002. PAUP*. Phylogenetic analysis using parsimony (*and other methods). Version 10. Sinauer Associates, Sunderland, Massachusetts. TABATADZE, E.S., AND V.A. YASNOSH. 1997. Control measures of Lopholeucaspis japonica Cockerell (Homoptera: Coccinea) through integrated citrus pest management. IOBC/WPRS Bulletin 20: 45-51. TABATADZE, E.S., AND V.A. YASNOSH. 1999. Population dynamics and biocontrol of the Japanese scale, Lopholeucaspis japonica (Cockerell) in Georgia. Entomologica 33: 429-434. TAKANO, Y., T. KIKUCHI, Y. KUBO, J.E. HAMER, K. MISE, AND I. FURUSAWA. 2000. The Colletotrichum lagenarium MAP kinase gene CMK1 regulates diverse aspects for fungal pathogenesis. Mol. Plant-Microbr. Interac. 13: 374-383. TALHINHAS, P., S. SREENIVASAPRASAD, J. NEVES-MARTINS, AND H. OLIVEIRA. 2002. Genetic and molecular characterization of Colletotrichum acutatum causing anthracnose of lupins. Phytopathology 92: 86-996. TALHINHAS, P., S. SREENIVASAPRASAD, J. NEVES-MARTINS, H. OLIVEIRA. 2005. Molecular and phenotypic analysis reveals the association of diverse Colletotrichum acutatum groups and a low level of C. gloeosporioides with olive anthracnose. Appl Environ. Microbiol. 71 (6): 2987-2998. TALERICO, R.L., C.W. MCCOMB, AND W.T. GARRETT. 1967. Fiorinia externa Ferris, a scale insect of hemlock.U. S. For. Serv. For. Pest Leafl. 107. 4 p.

189 TANADA, Y., AND H.K KAYA. 1993. Epizootiology. P. 595-632 in Insect Pathology, Tanada, Y. and H. K. Kaya (eds.). Academic Press, New York. TANAKA, M.A.S., AND F.A. PASSOS. 1998. Caracterização cultural e morfo-fisiológica de isolados de Colletotrichum causadores de antracnose do morangueiro em São Paulo. Summa Phytopathologica 24: 145-151. (In Portuguese) TANAKA, M.A.S., F.A. PASSOS, J.A. BETTI AND C.M.P. PIRES. 2001. Inoculation methods of Colletotrichum fragariae on strawberry. Scientia Agricola 58 (4): 725-729. (In Portuguese). THAXTER, R. 1914. Note on the ascosporic condition of the genus Aschersonia Montagne. Botanical Gazette 57 (4): 308-313. TAKAGI, S. 1963. Fiorinia externa on Tsuga sieboldii in Japan. Insecta Matsumurana 26: 115-117. TEBEEST, D.O. 1988. Additions to host range of Colletotrichum gloeosporioides f. sp. aeschynomene. Plant Disease 72 (1): 16-21. TEBEEST, D.O. 1991. Ecology and epidemiology of fungal plant pathogens studied as biological control agents of weeds. P. 97-114 in Microbial control of weeds. D.O. TeBeest (ed.). Routledge, Chapman & Hall, Inc. New York. TEBEEST, D.O., X.B. YANG, AND C.R. CISAR. 1992. The status of biological control of weeds with fungal pathogens. Annu. Rev. Phytopathol. 30: 637-657. TEBEEST, D.O., AND C.R. CISAR. 1994. Sexual cycle and potential for gene flow in fungal biological control agents. http://www.isb.vt.edu/brarg/brasym94/tebeest.htm (last access in 08/01/07) TEIXEIRA, M.A., W. BETTIOL, AND R. CESNIK. 2001. Patogenicity of Coletotrichum gloeosporioides, Orthezia praelonga pathogenic agent, to citrus leaves, flowers and fruits. Summa Phytopathologica 27: 352-357. (In Portuguese) TEIXEIRA, M.A., W. BETTIOL, R. CESNIK, AND R.F. VIEIRA. 2004. Pathogenicity of the fungus Colletotrichum gloeosporioides, pathogen of Orthezia praelonga, to several fruits and to pumpkin seedlings. Revista Brasileira de fruticultura 26 (2): 356-358. TEMPLETON, M.D., D.O. TEBEEST, AND R.J. SMITH JR. 1984. Biological weed control in rice with a strain of Colletotrichum gloeosporioides (Penz.) Sacc. used as a mycoherbicide. Crop Protection 3: 409-422. TEMPLETON, M.D., E.H.A. RIKKERINK, S.L. SOLON, AND R.N. CROWHURST. 1992a. Cloning and molecular characterization of the glyceraldehydes-3-phosphate dehydrogenase-encoding gene and cDNA from the plant pathogenic fungus Glomerella cingulata. Gene 122: 225-230 TEMPLETON, G.E. 1992b. Use of Colletotrichum strains as Mycoherbicides. P. 358-380 in Colletotrichum – Biology, pathology and control, Bailey, J. A. and M. J. Jeger (eds.). CAB International. Wallingford, UK. TENG, P.S. AND X.B. YANG. 1993. Biological impact and risk assessment in plant pathology. Annu. Rev. Phytopathol. 31: 495-521. TENHAKEN, R., A. LEVINE, L. F BRISSON, R. A., DIXON, AND C. LAMB. 1995. Function of the oxidative burst in hypersensitive disease resistance. PNAS 92: 4158-4163. THON M.R., AND D.J. ROYSE. 1999. Partial β-Tubulin gene sequences for evolutionoray studies in the Basidiomycota. Mycologia 91 (3): 468-474.

190 TIMMER, L.W., AND G.E. BROWN. 2000. Biology and control of anthracnose diseases of citrus. P. 300-316 in Colletotrichum – Host specificity, pathology and host-pathogen interaction. Prusky, D., S. Freeman and M.B. Dickman (eds.). APS Press. St. Paul, Minnesota. TSROR, L., O. ERLICH, AND M. HAZANOVSKY. 1999. Effect of Colletotrichum coccodes on potato yield, tuber quality, and stem colonization during spring and autumn. Plant Disease 83: 561-565. TU, J.C. 1980. The ontogeny of the sclerotia of Colletotrichum coccodes. Can. J. Bot. 58: 631-636. TU, J.C. 1992. Colletotrichum lindemuthianum on Bean: Population dynamics of the pathogen and breeding resistance. P. 203-224 in Colletotrichum – Biology, pathology and control, Bailey, J. A. and M. J. Jeger (eds.). CAB International. Wallingford, UK. TUBBS, C.H. 1995. Aspects of eastern hemlock silvics important in silviculture: an overview. P. 5-9 in Hemlock ecology and management. Proceedings of a regional conference on ecology and management of eastern hemlock. Mroz, G. and A.J. Martin (eds.). Dept. of Forestry, University of Wisconsin, Madison. TURGEON, B.G. 1998. Application of mating-type gene technology to problems in fungal biology. Ann. Rev. Phytopathol. 36: 115-137. UREÑA-PADILLA, A.R., S.J. MACKENZIE, B.W. BOWEN, AND D.E LEGARD. 2002. Etiology and population genetics of Colletotrichum spp. causing crown and fruit rot of strawberry. Phytopathology 92: 1245-1252. USDA FOREST SERVICE. 2004. Forest insects and diseases conditions in the United States 2003. USDA Forest Service Publications. 142 p. VAILLANCOURT, L., AND R.M. HANAU. 1994. Nitrate-nonutilizing mutants used to study heterokaryosis and vegetative compatibility in Glomerella graminicola (Colletotrichum graminicola). Experimental Mycology 18: 311-319. VAILLANCOURT, L., AND R.M. HANAU. 1999. Sexuality of self-sterile strains of Glomerella graminicola. Mycologia 91 (4): 593-596. VAILLANCOURT, L., J. WANG, AND R.M. HANAU. 2000. Genetic regulation of sexual compatibility in Glomerella graminicola. P. 29-44 in Colletotrichum – Host specificity, pathology and host-pathogen interaction, Prusky, D., S. Freeman and M.B. Dickman (eds.). APS Press. St. Paul, Minnesota. VAN EPENHUIJSEN, C. W., R.C. HENDERSON, A. CARPENTER, AND G.K. BURGE. 2000. The rise and fall of manuka blight scale: a review of the distribution of Eriococcus orariensis (Homoptera: Eriococcidae) in New Zealand. N. Z. Entomol. (23): 67-70. VENETTE, J.R., H.A LAMEY, AND L. LAMMPA. 1994. First report of lentil anthracnose (Colletotrichum truncatum) in the United States. Plant Disease 78: 1216. VERHOEFF, K. 1974. Latent infections by fungi. Ann. Rev. Phytopathol. 12: 99-110. VON ARX, J. A. 1957. Die arten der gattung Colletotrichum Cda. Phytopathol. Zeitschrift 29: 413-468. (In German) VIEGAS, E.C., H.N. SAMPAIO, P.O.L. CARVALHO, J.C. PERRUSO, AND P.C.R. CASSINO. 1995. Contole alternativo de Orthezia praelonga Douglas, 1891 (Homoptera, Ortheziidae) em laboratório. P. 333 in 15o Congresso de Entomologia, Caxambú, Brazil. (In Portuguese)

191 WALKER, J., A. NIKANDROW, AND G.D. MILLAR. 1991. Species of Colletotrichum on Xanthium (Asteraceae) with comments on some taxonomic and nomenclatural problems in Colletotrichum. Mycol. Res. 95 (10): 1175-1193. WALLER, J.M. 1992. Colletotrichum diseases of perennial and other cash crops. P. 167- 201 in Colletotrichum – Biology, pathology and control. Bailey, J.A. and M.J. Jeger (eds.). CAB International. Wallingford, UK. WALLNER, W.E. 1962. A field test with insecticides to control the scale Fiorinia externa on canadian hemlock. J. of Econ. Entomol. 55 (5): 798-799. WALLNER, W.E. 1964. Fiorinia hemlock scale – pest of ornamental hemlocks. Farm Res. 1 (30): 2-3. WALLNER, W.E. 1965. Biology and control of Fiorinia hemlock scale Fiorinia externa Ferris. PhD thesis, Cornell Univ., Ithaca, New York, 165 p. WALLNER, W.E. 1978. Scale insects: What the arboriculturist need to know about them. J. of Arboriculture 4 (5): 97-103. WAN, F.H., AND R. WANG. 2001. Biological weed control in China: An update report. P. 8-19 in Alien invasive species. Workshop on alien invasive species. Global diversity forum. South and southeast Asia session. Colombo, Sri Lanka. IUCN Regional biodiversity programme, Asia. IUCN – The World Conservation Union. WARWICK, S.I., AND L.D. BLACK. 1988. The biology of Canadian weeds. # 90. Abutilon theophrasti. Canadian J. of Plant Science 68: 1069-1085. WASILWA, L., J.C. CORRELL, AND T.E. MORELOCK. 1991. Relationship between vegetative (heterokaryon) compatibility groups and races of Colletotrichum orbiculare. Phytopathology 81 (10): 1191. (Abstr.) WATSON, J.R., AND E.W. BERGER. 1937. Citrus insects and their control. University of Florida, Agric. Exp. Bul. 88. 135 p. WATTS, P., P. KITTAKOOP, S. VEERANONDHA, S. WANASITH, R. THONGWICHIAN, P. SAISAHA, S. INTAMAS, AND N.L. HYWEL-JONES. 2003. Cytotoxicity against insect cells of entomopathogenic fungi of the genera Hypocrella (anamorph Aschersonia): possible agents for biological control. Mycol. Res. 107 (5): 581-586. WEBBER, H.J. 1894. Preliminary notice of a fungous parasite on Aleyrodes citri. R. and H. Journal of Mycology 7: 363-364. WEBBER, H.J. 1897. Sooty mold of the orange and its treatment. U.S. Department Agr. Div. Veg. Phys. and Path. Bul. 13: 1-44. WEI,Y.D., K.N. BYER, AND P.H GOODWIN. 1997. Hemibiotrophic infection of round- leaved mallow by Colletotrichum gloeosporioides f. sp. malvae in relation to leaf senescence and reducing reagents. Mycol. Res. 101 (3): 357-364. WEI, Y.D., W. SHEN, M. DAUK, F. WANG, G. SELVARAJN, AND J. ZOU. 2004. Targeted gene disruption of Glycerol-3-phosphate dehydrogenase in Colletotrichum gloeosporioides reveals evidence that glycerol is a significant transferred nutrient from host plant to fungal pathogen. Journal of Biological Chemistry 279 (1): 429- 435. WEISING K., F. WEIGAND, A.J. DRIESEL, G., KAHL, AND H. ZISCHER. 1989. Polymorphic simple GATA/GACA repeats in plant genomes. Nucleic. Acids Res. 17: 10128.

192 WEIDEMANN, G.J. 1991. Host-range Testing: Safety and Science. P. 83-96 in Microbial Control of Weeds. D.O. TeBeest (ed.). Routledge, Chapman & Hall, Inc. NY. WHITE, T.J., T. BRUNS, S. LEE, AND J.W. TAYLOR. 1990. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. P. 315-322 in PCR Protocols: A guide to methods and applications, Innis M.A., D.H. Gelfand, J.J. Sninsky and T.J. White (eds.). Academic Press, Inc., New York. WHEELER, H.E. 1950. Genetics of Glomerella VIII: A genetic basis for the occurrence of minus mutants. Am. J. Bot. 37 (4): 304-312. WHEELER, H.E. 1954. Genetics and evolution of heterotallism in Glomerella. Phytopathology 44: 342-345. WHITE T.J., T. BRUNS, S. LEE, AND J.W TAYLOR. 1990. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. P. 315-322 in A guide to methods and applications. PCR Protocols. Innis, M.A., D.H. Gelfand, J.J. Sninsky, T.J. White (eds.).Academic Press, Inc., NY. WILDING, N. 1983. The current status and potential of entomogenous fungi as agents of pest control. P. 743-750 in Proceedings of the 10th International Congress of Plant Protection. Brighton, England. WILLIAMS, D.W., M.E. MONTGOMERY, AND K.S. SHIELDS. 2002. Monitoring hemlock woolly adelgid and assessing its impacts in the Delaware river basin. P. 360-363 in Proceedings of the hemlock woolly adelgid in the eastern United States symposium, Onken, B., R. Reardon and J. Lashomb (eds.). New Jersey Agricultural Experiment Station, Rutgers University, New Brunswick, NJ. WILSON, M., L.V. MADDEN, AND M.A. ELLIS. 1992. Overwinter survival of Colletotrichum acutatum in infected strawberry fruit in Ohio. Plant Disease 76: 948- 950. WILSON, M., AND S.E. LINDOW. 1993. Release of recombinant microorganisms. Annu. Rev. Microbiol. 47: 913-944. WOLCOTT, N. 1940. Epidemics of fungus disease control insect pests in Puerto Rico. Scientific Notes 33 (1): 201-202. WRAIGHT S.P., M.A., JACKSON, AND S.L. DE KOCK. 2001. Production, stabilization and formulation of biocontrol agents. P. 253–288 in Fungi as biocontrol agents. Butt T.M., C. Jackson, N. Magan (eds.). CAB International. WYDEVEN, A.P., AND R.W. HAY. 1995. Mammals, amphibians and reptiles of hemlock forests in the Lake Superior region. P. 115-123 in Hemlock ecology and management. Proceedings of a regional conference on ecology and management of eastern hemlock. Mroz, G. and A.J. Martin (eds.). Dept. of Forestry, University of Wisconsin, Madison. WYMORE, L.A., AND A.K. WATSON. 1986. An adjuvant increases survival and efficacy of Colletotrichum coccodes, a mycoherbicide for velvetleaf (Abutilon theophrasti). Phytopathology 6 (10): 1115-1116. WYMORE, L.A., A.K. WATSON, AND A.R. GOTLIEB. 1987. Interaction between Colletotrichum coccodes and thidiazuron for control of Velvetleaf (Abutilon theophrasti). Weed Science 35: 377-383.

193 WYMORE, L.A., C. POIRIER, A.K. WATSON, AND A.R. GOTLIEB. 1988. Colletotrichum coccodes, a potential bioherbicide for control of velvetleaf (Abutilon theophrasti). Plant Disease 72: 534-538. XIAO, C.L., S.J. MACKENZIE AND D.E. LEGARD. 2004. Genetic and pathogenic analyses of Colletotrichum gloeosporioides isolates from strawberry and noncultivated hosts. Phytopathology 94: 446-453. YAKOBY, N., R. ZHOU, I. KOBILER, A. DINOOR, AND D. PRUSKY. 2001. Development of Colletotrichum gloeosporioides restriction enzyme-mediated integration mutants as biocontrol agents against anthracnose disease in avocado fruits. Phytopathology 91: 143-148. YAKOBY, N., I. KOBILER, A. DINOOR, AND D. PRUSKY. 2000. pH regulation of pectate lyase secretion modulates the attack of Colletotrichum gloeosporioides on avocado fruits. Appl. and Environ. Microbiology 66 (3): 1026-1030. YANG, X.B., AND D.O. TEBEEST. 1992. Green treefrogs as vectors of Colletotrichum gloeosporioides. Plant Disease 76 (12): 1266-1269. YANG, X.B., AND D.O. TEBEEST. 1993. Epidemiological mechanisms of mycoherbicide effectiveness. Phytopathology 83 (9): 891-893. YANG, X.B., AND D.O. TEBEEST. 1994. Distribution and grasshopper transmission of northern Jointvetch anthracnose in rice. Plant Disease 78 (2): 130-133. YASNOSH, V.A., AND E.S. TABATADZE. 1997. Aschersonia fungus (Deuteromycetes). A new entomopathogens of armored scale insect in Georgia. Mikologia and Fitopatologia 31: 59-63. (In Russian) YORKS, T.E. 2002. Influence of hemlock mortality on soil water chemistry and ground flora. P. 47-49 in Proceedings of the hemlock woolly adelgid in the eastern United States symposium. Onken, B., R. Reardon and J. Lashomb (eds.). New Jersey Agricultural Experiment Station, Rutgers University, New Brunswick, NJ. ZHU-AN, C., AND C. QING-TAO. 1986. An entomogenous fungus of the whitefly Aleurotrachelus camellia Kuwana. Acta Mycologica Sinica 5 (1): 37-43. (In Chinese) ZHU, P., AND P. V. OUDEMANS. (2000). A retrotransposon Cgret from the phytopathogenic fungus Colletotrichum. Curr. Gen. 38: 241-247. ZHUAN, C. 1988. Some epidemiological characteristics of entomogenous fungi Aschersonia duplex Berk. and its application. Acta Phytophylacica Sinica 15 (4): 235- 239. (In Chinese) ZIMMERMANN, G. 1986. Insect pathogenic fungi as pest control agents in Jost M. Franz (ed.). Biological plant health protection. Fortschr. Zool. 32: 217-232. ZULFIQAR, M., R.H. BRLANSKY, AND L.W. TIMMER. 1996. Infection of flower and vegetative tissues of citrus by Colletotrichum acutatum and C. gloeosporioides. Mycologia 88 (1): 121-128.

194 APPENDICES

Appendix 1. The genus Aschersonia

The Hyphomycete genus Aschersonia, named by Montagne (1848) is the conidial stage of the teleomorph Hypocrella (Clavicipitaceae). Most of known species occur in the

Aschersonia form. Perithecia are rarely collected (Mains, 1959a). The great majority of

species were initially described as having a pantropic distribution (Thaxter, 1914).

Hypocrella spp. were first reported in Cuba, Puerto Rico, Paraguay and the Philipines in the leaf surface of different plant species in the early 20st Century (Seaver, 1920).

Although the genus was described as parasitic in plants, it was later established that it is

parasitic on insects in the Hemipteran families Coccidae (soft scales), Diaspididae

(armored scales) and Aleyrodidae (whiteflies), infesting the plants where they are found

(Petch, 1921). Species of this genus have been used as a biological control for Japanese

citrus scale, Lopholeucaspis japonica (Cockerell) in the Black Sea coast of Georgia

(Tabatadze and Yasnosh, 1997; Yasnosh and Tabatadze, 1997; Tabatadze and Yasnosh,

1999).

Until 1919, 60 species had been described (Petch, 1921). However, only 54

scientific names covering 42 species were valid. In the species group parasitic to scale

insects, Petch (1921) stated there were 22 species of Hypocrella, with the corresponding

Aschersonia anamorph being known in 12 cases. Three unattached species of

Aschersonia (teleomorph not known) where also listed. For whiteflies, there were 13 species of Hypocrella of which anamorph Aschersonia stages were known, and 14 unattached species of Aschersonia (Steinhaus, 1949). Petch (1925) was able to examine a significant portion of all herbarium material in existence around the world and estimated

195 that there were 25 Aschersonia spp. and 29 Hypocrella spp. Apart form the work developed by Petch, few taxonomical studies have been conducted, with the exception of

Mains (1959b) who reviewed the number of Aschersonia species, reporting the existence of seven Aschersonia spp. occurring in North America. Currently, this genus is lacking an extensive taxonomical revision and Evans and Hywel-Jones (1997) noted that many species from tropical forest ecosystems remain to be described.

Members of Aschersonia have a superficial stroma (when present, on the lower surface of the leaves), no hyphae penetrating the leaf, generally white or brightly colored in various shades of yellow, red or brown turning black with age in most species, and a subglobose, hemispherical or pulvinate shape (Petch, 1914, 1921). The conidia spores are usually ellipsoid, granular and not septated, although variations in the shape have been reported. Spores are born singly on short filiform hyphae (Samson and McCoy, 1983). In early stages of infection, the fungus develops internally within the host and thus not visible externally. Soon, a dense marginal fringe is formed by the hyphae bursting out from the interior around the edge of the host (Fawcett, 1948). In its mature stage the fungus completely covers the insect which in some cases makes it difficult to identify the parasitized hosts. Cross sections of the stroma may not show recognizable remnants of the host (Samson and McCoy, 1983), hence the past assumption that the fungus was parasitic on leaf tissue.

Webber (1894, 1897) first reported the entomogenous condition of the anamorph

Aschersonia while conducting investigations in entomopathogenic fungi on citrus trees.

Gossard (1903) and Fawcett (1948) continued Webber’s work. Since 1904, species of

Aschersonia have been described as occurring on scale insects (Petch, 1921), especially

196 in the tropics, but also present in temperate regions (Australia, Africa, India, Java, New

Zealand and North America) attacking the insects in epizootic patterns (Parkin, 1906).

The successful use and establishment of several Aschersonia spp. (mainly

Aschersonia aleyrodis Webb.; Aschersonia goldiana Sacc. and Ellis and Aschersonia placenta Berk. et Br.) for control of greenhouse whiteflies (Hemiptera: Aleyrodidae) has been reported in North America (Fawcett, 1908a,b, 1948; Berger, 1919, 1921, 1944;

McCoy, 1985), Germany (Hirte et al., 1989), The Netherlands (Fransen 1987, 1995,

Fransen and Van Lenteren, 1993, 1994, Meekes et al. 2000, 2002), eastern Europe

(Ponomarenko et al., 1975; Solovei, 1976; Spassova, 1980; Shutova et al., 1985),

Colombia (Sarmiento et al., 1995) and Asia (Zhu-An and Qing-Tao, 1986). As a result of the successful biocontrol achieved with Aschersonia spp. in citrus pests, its use has been extended to cucumber and tomato whitefly pests (Spassova et al., 1980; Ramakers and

Samson, 1984). The toxigenic properties of metabolites from Aschersonia spp. have also been studied (Krasnoff and Gibson, 1996; Boonphong et al., 2001, Watts et al., 2003).

Aschersonia spp. have been extensively studied for the biological control of whiteflies.

The potential use of Aschersonia for management of scale insects has been neglected considering the relevant importance of some species of Diaspidids. Research within this area urges since to accomplish effectiveness, several factors must be equated, including assessing potential, efficacy, dissemination, persistence, compatibility with other regulatory agents, and establishment in the host population (see Fuxa, 1987; Jaques,

1983).

197