BIOLOGICAL CONTROL OF COGONGRASS. Imperata cylindrica (L.) Beauv.

CAMILLA B. YANDOC

A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTL\L FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

. ^ 2001 . Copyright 2001

by

Camilla B. Yandoc To my parents, Francisco and Marietta Yandoc ACKNOWLEDGMENTS

I would like to thank my major professor, Dr. Raghavan Charudattan, for

believing in me and for giving me the opportunity to enrich myself both professionally

and personally. I also would like to thank Dr. Francis W. Zettler and my committee

members, Dr. Richard D. Berger, Dr. James W. Kimbrough, and Dr. Donn G. Shilling,

for the time, interest and support they have given me. Many thanks go to Jim DeValerio,

Karen Harris, Jay Gideon, Henry Ross, Jason McCombs, Matt Pusateri, Josh Rhames,

Mark Elliott, Eldon Phihnan and Herman Brown for all the help they provided during the

course of my experiments and for their friendship. Thanks go to my former labmates. Dr.

Erin Rosskopf, Dr. Jugah Kadir, Dr. S. Chandramohan, Dr. Dauri Tessman, Dr. Gabriela

Wyss, and Ms. Angela Vincent. Thanks go to all my friends in the Plant Pathology

Department, Mr. Lucious Mitchell, Mr. Gene Crawford, the USPS, the graduate students,

Ms. Lauretta Rhames, Ms. Donna Belcher, Kendra Davis, and Ms. Gail Harris, and

everyone else who helped me in one way or another. Many thanks go to Dr. Ziyad

Mahfoud and Ms. Upasana Santra for helping me with the data analysis.

I would like to thank my family, my Mom and Dad, my sisters. Carmen and

Guadalupe, brother Deodato, my nephews Zygie and Mickey, and my niece Miranda for

all the love and support. Thanks go to my niece Isabella, for reminding me to always count my blessings. Thanks go to Gino, for his love, friendship and patience. Thanks go to Antonio and Sheila Gomez and family. Bob and Pam Kemerait, Dr. and Mrs. Abies

. iv and, and to all my friends back home and all my friends in Gainesville. Finally, I thank

God for always making things work out fine.

V TABLE OF CONTENTS

ACKNOWLEDGMENTS iv

ABSTRACT ix

CHAPTERS U-

1 INTRODUCTION .1 . : 1

Cogongrass Introduction into the United States 7 Economic and Ecological Importance of Cogongrass 8 Morphology and Biology 12 Habitat 14 Cogongrass Management 15

2 EVALUATION OF FUNGAL PATHOGENS AS BIOLOGICAL CONTROL AGENTS FOR COGONGRASS 23

- Introduction 23 Materials and Methods 25 Results 30 Discussion 37

3 MASS PRODUCTION OF INOCULUM OF Bipolaris sacchari AND Z)recA5/era g/^an/ea " AND DETERMINATION OF . ' OPTIMUM CONDITIONS TO MAINTAIN SPORE " VL\BILITY DURING STORAGE 44

i ^ Introduction 44 Materials and Methods 49

vi ,

Results 54 Discussion 65

4 THE EFFECT OF OIL FORMULATION ON THE BIOHERBICIDAL EFFICACY OF Bipolaris sacchari AND Drechslera gigantea 70

Introduction 70 Materials and Methods 73 Results 79 Discussion 93

5 EVALUATION OF HOST RANGE OF Bipolaris sacchari AND Drechslera gigantea 97

Introduction 97 Materials and Methods 100 Results 102 Discussion 103

6 FIELD EFFICACY OF Bipolaris sacchari and Drechslera gigantea AS BIOHERBICIDES OF COGONGRASS Ill

Introduction Ill Materials and Methods 116 Results 120 Discussion 136

7 SUPPRESSION OF COGONGRASS WITH BIOHERBICIDE APPLICATION AND PLANT COMPETITION 142

Introduction 142 Materials and Methods 144 Results 148 Discussion 158

vii .1

8 SUMMARY AND CONLUSIONS 1 62

REFERENCES. .168

BIOGRAPHICAL SKETCH 1 94 : \ Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy

. BIOLOGICAL CONTROL OF COGONGRASS . . w

Camilla B. Yandoc

December, 2001

Chairman: Raghavan Charudattan Major Department: Plant Pathology

Two fungal pathogens, Bipolaris sacchari and Drechslera gigantea, were evaluated for their potential as biological control agents for cogongrass (Imperata cylindrica [L.] Beauv.). Application of spore suspensions of each of these fungi that contained lO'* -10^ spores/ml and 1% gelatin resulted in 30-50% disease severity, on a rating scale with 50% as the maximum disease severity. Disease symptoms ranged from foliar lesions to blighting. Spores formulated in an oil emulsion caused more disease severity in the greenhouse trials. Mass production of spores was achieved in the laboratory with sohd substrate culturing techniques. Spores dried at 30-40''C for 5 days and stored at low temperature (10-25°C) and low humidity (30-60%) RH) remained viable for at least 10 weeks. In host-range tests, 19 plant species in six families (Alliaceae,

Apiaceae, Asteraceae, Cucurbitaceae, Poaceae, and Solanaceae) were found to be

ix susceptible to B. sacchari formulated in oil emulsion. Susceptibility of the species outside the family Poaceae was attributed to the predisposition of the test plants by the oil formulation and their age and vigor. ^ '

Among the resistant grass species was bahiagrass, a commonly used turf and

experiments in the greenhouse, B. sacchari pasture grass species in Florida. In | selectively controlled cogongrass and reduced its ability to compete with bahiagrass in cogongrass-bahiagrass mixtures. Thus, B. sacchari can be used to suppress cogongrass and to allow the establishment of a more desirable species, such as bahiagrass.

In field trials, there was a significant interaction of the spore concentration, oil concentration of the carrier, and application volume on the efficacy of B. sacchari and D. gigantea. Generally, application of a larger volume of inoculum (800 ml/m^) that contained 10^ or 10^ spores/ml and 18 or 26% oil, applied with a hand-pumped sprayer, resulted in the highest levels of foliar blighting (>70% on a rating scale of 0-100%) during the 6-week experimental period. Application of 5. sacchari and D. gigantea, singly or as a mixture at the rate of 200 ml/m with 10 spores/ml with an ultra- low- volume-applicator resulted in >70% disease severity in the field. Bipolaris sacchari and

D. gigantea had the efficacy to be biological control agents of cogongrass. CHAPTER 1

INTRODUCTION

Human activity has greatly influenced the movement of plant species worldwide.

Through man, plants are able to overcome natural barriers to dispersal and reach new geographical sites that they could not otherwise reach. Plant explorations carried out by public agencies, university researchers, plant collectors, botanical gardens, and people in the terrestrial and aquatic ornamental plant trade have brought about the deliberate and

the accidental release of non-native plant species (Simberloff et al., 1997). Accidental introductions occurred when non-native plants enter as contaminants in agricultural seed, packing material, shipping containers, or ballast or on vehicles and other conveyances

(Office of Technology Assessment, 1993). Through the years, the deliberate and accidental introductions of plant species have accelerated with the increase in transportation and international commerce (Cronk and Fuller, 1995; Randall and

MarinelU, 1996; Wagner, 1993).

* By 1923, there were about 50,000 non-native plant species that have been introduced into the United States. Most of these non-native plant species were brought in for food, fiber, medicine, and ornamental purposes or for scientific curiosity. While most non-native plants have been beneficial to society and are ecologically benign, some of the introduced plant species have caused major economic losses in agriculture, forestry, pastures, and several other sectors of the U.S. economy, in addition to bringing harm to the environment (Babbitt, 1998; Pimentel et al., 1999). Approximately 500 introduced species have become serious weeds in crop systems (Pimentel et al., 1989) and about

5,000 introduced plant species have escaped and now exist in natural ecosystems, as compared to about 17,000 species of native plants (Morin, 1995; Morse et al., 1995).

Non-native plant species that have grown out of their areas of initial introduction and have encroached into the habitats of indigenous plant populations are considered invasive weeds (Westbrooks, 1998). The absence of natural enemies, which limit plant reproduction and spread, has allowed a number of introduced non-native species to take over natural areas and disrupt native plant communities (Randall and Marinelli, 1996).

The natural enemies may include vertebrate and invertebrate herbivores, plant pathogens, and competing plants. Human and natural disturbances (i.e., fire, grazing, soil disturbance, trampling, and habitat fragmentation) that remove native vegetation also allow for the establishment of invasive species in natural communities (Hobbs and

Huenneke, 1992; McEvoy and Rudd, 1993).

Invasive weeds are capable of changing the composition, structure, or fimction of the ecosystem they invade (Cronk and Fuller, 1995). Next to habitat destruction and direct human exploitation of organisms, invasive species are considered as the second most important threat to biodiversity (Randall, 1996; Pimm and Gilpin, 1989). Invasive plant species can alter ecosystem processes, displace native plant and animal species, support non-native animals and microbes, and hybridize with native species, and subsequently alter gene pools (Randall, 1996). ^ .

Invasive plant species threaten biodiversity by suppression of native plant populations through competition or allelopathy and thus, modify the habitat of native animals. There are a number of characteristics that permit invasive plants to take over new areas rapidly and out-compete native plants for Ught, water, and nutrients. Some of these features include: early maturation, profuse reproduction by seeds or by vegetative structures, seed dormancy, adaptations for successful dissemination, production of toxins that suppress other plants, morphological features that deter herbivory, roots or rhizomes with large food reserves, survival and seed production under adverse environmental conditions, and high photosynthetic rates (Westbrooks, 1998). Several non-native grass species, in particular, are successful invaders of natural communities because they can

- tolerate or enhance fire, or respond to fire with rapid growth (D'Antonio and Vitousek,

1992).

Non-native weeds are spreading and invading about 700,000 ha of U.S. wildlife habitat annually (Babbitt, 1998). The European purple loosestrife (Lythrum salicaria

L.), introduced as an ornamental in the early 19* century (Malecki et al., 1993), is

spreading at the rate of 1 1 5,000 ha/yr and is changing the basic structure of most

wetlands it has invaded (Thompson et al., 1987). The European cheatgrass (Bromus

tectorum L.) is another example of an invasive weed that is dramatically changing the vegetation and fauna of many natural ecosystems and predisposing the invaded habitats

to fires (Kurdila, 1995; Vitousek et al., 1996; Vitousek et al., 1997). Cheatgrass now occurs on 5 million ha in Idaho and Utah (Whisenant, 1990). In Florida, more than 900 of the 25,000 ahen plant species imported as ornamentals have escaped and have become

established in surrounding natural ecosystems (Frank and McCoy, 1995; Frank et al.,

1997; Simberloff et al., 1997). The Australian meleleuca (Meleleuca quinquenervia

[Cav.] S.T. Blake), introduced into Florida for ornamental and agricultural purposes 4 . j.,-.;.,,

(Crowder, 1974), is invading wetland ecosystems at the rate of 2,850 ha per year or 7.8

ha per day (Center et al., 2000). Hydrilla {Hydrilla verticillata [L.f.] Royle) and water hyacinth {Eichhomia crassipes [Mart.] Solms), which were introduced into Florida as ornamental aquatic plants, now affect fish and other aquatic species, choke waterways, alter nutrient cycles, and reduce recreational use of water bodies (Center et al., 1997).

The economic impact of non-indigenous weeds consists of lost revenues (losses) and costs. In the agricultural sector, losses result from yield and quality reduction caused by weeds. Overall, weeds cause about 12% reduction in U.S. crop yields. This reduction translates to $33 billion in lost crop production annually, based on the potential value of

all U.S. crops of more than $267 billion/yr (United States Bureau of the Census, 1998).

Non-native weeds, which comprise about 73% of the major weed species in all of U.S. crops (Pimentel, 1993), are said to be responsible for about $24 billion/yr in crop losses

(Pimentel et al., 1999). Approximately $500 million is spent on the control of non-native weeds in residential areas while $1 billion is spent on the control of non-native weed

species in golf courses (Pimentel et al., 1999). A total of $100 million is invested annually in the United States for the control of non-indigenous aquatic species (Office of

Technology Assessment, 1993). In Florida alone, $19.3, $8.84, and $13 million were spent in 1999-2000 by the Florida Department of Environmental Protection (DEP), the

South Florida Water Management District (SFWMD), and the Florida Department of

Transportation (FDOT), respectively, for the management of invasive terrestrial and aquatic plant species occurring in waterways, roadsides, rights-of-way, forests, and other natural areas (Florida Department of Environmental Protection, 2000). Inputs or costs accrue as a result of herbicide use and the employment of tillage, mowing, and cultural and biological inputs for weed management and control (Bridges,

1999). In the 1980s, farmers spent over $3 billion annually for chemical weed control

and approximately $2.6 billion for cultural, ecological, and biological control methods

(Ross and Lembi, 1983). About 70-80% of pesticides used in the United States were

herbicides (Pimentel, 1991; Radosevich et al., 1997). hi 1997, the cost of herbicides used

by all sectors in the United States (agricultural, industrial, commercial, government, and

home and garden) amounted to about $6.8 billion (Aspelin and Grube, 1999).

The environmental impact of non-indigenous weeds results from the invasion of

native plant and animal habitats. Another environmental cost of weeds is the impact

associated with weed control agents and practices, such as herbicides, cultivation, tillage,

and fire. The use of herbicides may result in the injury of nontarget plant species and the

contamination of ground and surface water. Cultivation may result in surface run-off, silt

and sediment production, and possibly increased herbicide deposition in streams and

lakes due to herbicides that are adsorbed to mineral and organic soil fractions that are

transported from fields (Baker, 1999). Tillage predisposes the soil to wind and water

erosion (Baker, 1999; Wicks et al., 1995).

The continued use and over-use of herbicides has lead to the widespread

occurrence of herbicide resistance and cross-resistance in some weed species (Baker and

Terry, 1991). Triazine-resistant weeds in the United States were first reported in 1970

(Ryan, 1970). In the 1990s, at least 100 species were reported to have developed

herbicide-resistant biotypes. Resistance has been reported for triazines and 15 other

herbicides or herbicide families (LeBaron and McFarland, 1990). Resistance of some

• I 6

biotypes to newer target-specific herbicides, such as imidazoUnones and sulfonylureas, has also been reported (Primiani et al, 1990; Mallory-Smith et al., 1990). Evidence is mounting that some biotypes resistant to imidazoUnones may also be resistant to

sulfonylureas, i. e., they are as cross-resistant. Cross-resistance to herbicides with dissimilar modes of action has also been reported (Powles and Howat, 1990).

Plant pathogens, as well as insects have been considered as potential cost- effective means to reduce weed populations in crop systems, forests, rangelands, and even in natural areas where the low-profit margins preclude large herbicidal expenditures

(TeBeest et al., 1992). The study and utilization of parasites, predators, and pathogens for the regulation of host population densities is termed biological control (DeBach,

1964). Biological control of weeds with plant pathogens may be employed by the classical, the augmentative, or the inundative (bioherbicide) approach. The classical approach consists of a single or a limited number of inoculative introduction of a biological control agent into a region where the target weed has become a newly invaded pest (Hokkanen, 1985; Waage and Greathead, 1988). The augmentative approach consists of periodic releases of the control agent to augment an existing population of a

biocontrol agent that is not sufficient enough to suppress the target weed population

(Harley and Fomo, 1992). The inundative approach involves the application of the biocontrol agent in massive doses to create an epidemic within the target weed population

(Templeton et al., 1979; Charudattan, 1984).

Plant pathogens employed as biological control agents with the inundative approach are called bioherbicides. Fungal pathogens that are formulated as bioherbicides are specifically referred to as mycoherbicides. Two aspects, namely, the in vitro culture 7

and mass production of fungal inoculum, and the inundative application of the inoculum

distinguish the to achieve a rapid epidemic buildup and a high level of disease, mycoherbicide concept (Charudattan, 1988). Bioherbicides offer similar advantages that chemical herbicides have over mechanical control methods, which include host selectivity and reduced erosion risk. Bioherbicides do not pose the threat of non-target

pollution injury from drift or leaching. They are biodegradable, and result in negligible problems (Auld and Morin, 1995). The use of bioherbicides is a viable alternative

method for control of specific weeds or situations for which no suitable chemical

herbicides are available, such as lakes, waterways, rangelands, forests and other natural

areas infested with weeds. Bioherbicides are also valuable in low-value crops and minor

crops that lack registered herbicides.

Cogongrass Introduction into the United States

The accidental introduction of cogongrass (Imperata cylindrica [L.] Beauv.) from

the Old World to the United States occurred in 1911, through a shipment of Satsuma

orange from Japan to Mobile County, Alabama (Tabor, 1952; Dickens, 1974).

Cogongrass then reached Mississippi and Florida through a series of intentional

introductions between the 1920s and the 1940s (Tabor, 1949). Cogongrass from the

Philippines was infroduced into Florida at Gainesville, Brooksville, and Withlacoochee

for forage and soil stabilization purposes. By 1949, cogongrass had spread to northwest

Florida, Central Florida, and other portions of the state due to unauthorized plantings by

cattlemen (Tabor, 1949; Dickens and Buchanan, 1971; Dickens, 1973). The inadvertent

fransport in forage and soil during roadway construction and routine maintenance with

fill soil contaminated with cogongrass rhizomes also contributed to the continuous spread 8

of cogongrass (Bryson and Carter, 1993; Colvin et al., 1994). At present, cogongrass occurs as a weed in Alabama, Florida, Georgia, Louisiana, Mississippi, South Carolina,

Texas, and Virginia, and it continues to spread (Byrd and Bryson, 1999).

Small (1933) reported the occurrence of Imperata brasiliensis Trin. also known as the Brazilian satintail, Brazilian bladygrass, or silver plume, in the United States. It was

first collected from Miami-Dade, Florida in 1905 (Westbrooks and Eplee, 1989).

Imperata brasiliensis occurs in Mexico, South America, West Indies, and southern

Florida. This species is also naturalized around Mobile, Alabama and in Louisiana (Hall,

1998). According to McDonald et al. (1996), the crosses of/, cylindrica and/.

brasiliensis produced viable seeds. Both /. cylindrica and / brasiliensis are considered forms of the same species because of the frequent evidence of hybridization (Hall, 1998).

The cogongrass variety Red Baron or Japanese Blood Grass {Imperata cylindrica var. rubra), which is sold as an ornamental grass in the northeast, may also hybridize with /

cylindrica (Shilling et al., 1997). Unlike cogongrass, which has not shown tolerance to

low temperature, the Red Baron or Japanese Blood Grass is very freeze-hardy.

Infroduction of the Red Baron in the southeastern United States and its hybridization with cogongrass could produce biotypes and extend the range of cogongrass northward and westward (Bryson and Carter, 1993; Shilling et al, 1997).

Economic and Ecological Importance of Cogongrass

Cogongrass is the seventh worst weed of the world and the most serious perennial

weed of southern and eastern Asia (Holm et al., 1977). It infests over 500 million ha worldwide, 200 million ha in Asia, and several thousand ha in the southeastern United States (Dickens, 1974; Holm et al., 1977; Falvey, 1981). It is a weed of 35 economically important crops in 73 countries in the tropical regions of the world. Cogongrass is a serious weed of banana (Musa paradisiaca L.var), citrus {Citrus spp.), coconut {Cocos nucifera L.), oil palm (Elaeis guineensis Jacq.), pineapple {Ananas comosus (L.) Merr.), pastures, pine {Pinus spp.), rubber {Hevea brasiliensis [Willd. Ex A. Juss.] Mull. Arg.), tea {Camellia sinensis [L.] Kuntze), com {Zea mays L.), cassava {Manihot esculenta

Crantz), peanuts {Arachis hypogaea L.), upland rice {Oryza sativa L.), cotton {Gossypium spp.), sugarcane {Saccharum officinarum L), soybean {Glycine max [L.] Merr.), coffee

{Coffea spp.), and various vegetable crops (Holm et al., 1977). Cogongrass has the . potential to be a serious weed in the southeastern United States, particularly in agronomic crops grown with minimum tillage (Patterson et al, 1980).

production, Cogongrass has little utility except for thatch, short-term forage , erosion control, and in some areas, paper making (Watson and Dallwitz, 1992). It is an inferior forage crop for domesticated animals. The silica bodies throughout the leaves

and the razor-sharp leaf margins render it undesirable and unpalatable to grazing animals

(Coile and Shilling, 1993). The herbage yields of cogongrass are relatively low even under heavy fertilization. The nitrogen, phosphorous, and energy contents are very low.

The crude protein level is not enough to sustain cattle (Colvin et al., 1994). ' <

As early as 1948, a call was made for the eradication of cogongrass in the

Western Hemisphere because its weedy habit outweighed its usefulness as a forage crop

(Pendleton, 1948). The weedy habit of cogongrass also makes it an unsuitable species for

soil stabilization. It frequently spreads over large areas and excludes other grasses by

forming dense mats of branched rhizomes. The sharp apical tips of its rhizomes can 10

penetrate and injure the roots of crop plants (Boonitee and Ritdhit, 1984; Eussen and

Soerjani, 1975), and microorganisms may enter these injured roots and affect the other parts of the crops (Tempany, 1951; MacLagan Gorrie, 1950). Cogongrass also makes the

establishment of other desirable perennial grass species difficult because it can extract

soil moisture fi-om the shallow layers of the soil. It is able to invade areas that will not

support the growth of other grasses due to its ability to tolerate a wide range of soil

conditions (Hubbard et al., 1944). Cogongrass rhizomes exude allelopathic substances

that inhibit other plant species (Hussain et al., 1992). Allelopathy has a significant role in

the interference of cogongrass against other plant species (Rice, 1995). Leachates of

cogongrass leaves, roots, and rhizomes adversely affect mustard, fenugreek, and tomato

by reduction in seed germination, root growth, shoot growth, and fi-esh and dry weight.

Cogongrass also affected fungi in the soil, the number of colonies of Aspergillus fumigatus Fresen, A. niger Tiegh, and A. candidus Link, per gram of soil decreased while

the number of colonies of A. flavus Link: Fr. per gram of soil increased in soils where

cogongrass was present (Inderjit and Dakshini, 1991). Cogongrass is highly pyrogenic

and burning stimulates its vegetative spread. According to Lippincott (1997), in a

sandhill ecosystem where low-intensity fires fi-equently occur, the presence of cogongrass

can produce hotter, widespread, and frequent fires that can seriously damage or kill most

fire intolerant species. The favorable effect of fire on cogongrass allows it to be a

dominant species in this kind of ecosystem. Because it can alter the fire regime,

cogongrass has the potential to form grasslands that are devoid of trees and prone to fires

(Lippincott, 1997). 11

The USDA Animal and Plant Health Inspection Service (APHIS), Plant

Protection and Quarantine, and the Florida Department of Agriculture and Consumer

Services, Division of Plant Industry have listed cogongrass as a noxious weed (Coile and

Shilling, 1993). Cogongrass is included in the 1993 Florida Exotic Pest Plant Council

(EPPC) List of Most Invasive Plant Species. In Florida, cogongrass thrives in natural and disturbed areas such as grasslands, roadsides, forests, recreational areas, and reclaimed

• mine areas (Coile and Shilling, 1 993). The Florida Department of Transportation .

(FDOT) Bureau of Maintenance cited cogongrass as one of the nine grassy weed species that require additional maintenance efforts to ensure safe and aesthetically pleasing

roadways (Willard et al., 1990). According to an FDOT report, there are 900 large

monotypic stands of cogongrass, occupying more than 741 ha of right-of-way in Florida

(Caster, 1994). Aside from occupation of roadsides and utility corridors, cogongrass also

occupies natural areas in over 20 coimties. According to the Florida Department of

Forestry, most of the Withlacoochee State Forest (Citrus, Hernando, Sumter, and Pasco

counties) may be infested with cogongrass within a generation (Hardin, 1994). It is

considered a threat to nine state forests, that include habitats of federally listed

endangered species and threatened native plant species (Langeland and Burks, 1998).

Cogongrass is a poor habitat for native wildlife in Florida. According to

anecdotal information, gopher tortoises (Gopherus polyphemus [Daudin.]) abandon areas

infested with cogongrass. The loss of the gopher tortoise results in the disappearance of

numerous species that depend on gopher tortoise burrows, including gopher frogs (Rana

capita aesopus Cope), eastern indigo snakes {Drymarchon corais Boie), and scarab

beetles (Scarabaeidae). Cogongrass may also affect the nesting and foraging of the 12

endangered red-cockaded woodpecker (Picoides borealis Vieillot) by inhibition of regeneration of long-leaf pine. The trees in which woodpeckers nest are vulnerable to intense fires that can occur in cogongrass ground cover (Myers, 1990). Leaves of cogongrass persist after winterkill and are considered as a fire hazard especially when mixed with other volatile fiiels such as waxmyrtle, gallberry, and young pines (Shilling et

al., 1995).

Morphology and Biology

Cogongrass grows in loose or compact tufts with slender, erect leaves that arise

fi-om creeping rhizomes. Rhizomes are extensive, long and tough, with sharp-pointed

growing tips. They possess non-green leaves, called cataphylls, which give them a scaly

appearance. Cogongrass plants are virtually stemless, with one to eight leaves that

originate just above the ground. Plants are usually 1 m (3 ft) tall but may grow as high as

3 m (9 ft). The leaves are 4 to 10 mm wide and up to 15 to 150 cm long, with a whitish,

off-center midrib. Leaf margins are scarious (translucent and dry) and are scabrous

(rough). The spikelets are grouped into a panicle. The panicles are spike-like, cylindrical,

and dense. They are 3 to 20 cm long, 0.5 to 2.5 cm wide and are silky in appearance.

Each spikelet has a plume-like structure that allows the seeds to be carried by the wind.

Cogongrass reproduces both by seeds and rhizomes, ft is a prolific seed producer,

capable of producing 3,000 seeds per plant (Holm et al., 1977; McDonald et al., 1996).

The seeds are easily carried by the wind and can travel over long distances; the

aerodynamic properties of the spikelet clump allow seed dispersal to occur up to 24 km

over open country (Hubbard et al., 1944). Wind dispersal of seeds plays a significant role -

in the distribution of cogongrass (Dickens, 1974). In Asia, the spread of cogongrass from the coastal areas inland was facilitated by seed dispersal, primarily along the rights-of way bordering highways and railways (Hubbard et al, 1944). In Alabama, the spread of cogongrass northward appears to have occurred via wind dispersal of seeds on the northeasteriy winds off the Gulf of Mexico along Interstate 65 (Shilling et al., 1995;

Wilcutetal., 1988). V - H < ' f ,

Cogongrass flowers along the Gulf Coast area of Alabama, Mississippi, and

Florida from March to May, occasionally flowering in the early fall following a frost

(Dickens and Moore, 1974). Willard (1988) reported that cogongrass might flower all

year round in cenfral and south Florida. Flowering is induced by sfress due to

defoliation, mowing, grazing, and cool temperatures, and nifrogen fertilization (Holm et

al., 1977; Coile and Shilling, 1993). ^

Cogongrass seeds germinate within the range of 20°C to 35°C, with 30°C as the

optimum constant temperature. Germination is increased and hastened by exposure to

light. The seeds do not have a dormancy period and mature seeds germinate within a few

days after harvest (Dickens and Moore, 1974). Sexual reproduction through seeds allows

for the development of ecotypes adapted to new environments or more resistant to

management strategies (Shilling et al., 1997). . f

Asexual reproduction in cogongrass is achieved through rhizome production,

which allows cogongrass to spread rapidly over newly colonized areas. Plants grown

from rhizome fragments could produce up to 168 rhizomes in 87 days. Plants grown

from seeds can produce rhizomes within 30 to 40 days (Patterson et al., 1979). It was

estimated that there could be 6 metric tons of rhizomes in a hectare of land (Soerjani, 14

1970). Both roots and rhizomes possess structures that enable them to conserve water and resist desiccation, breakage, and fire (Holm et al., 1977). The roots may penetrate to

150 cm and the rhizomes may penetrate as deep as 120 cm. Rhizome development is enhanced by exposure to full sunlight, high temperatures, and long photoperiods

(Patterson et al., 1980; Wilcut et al., 1988).

Like most of the world's worst weeds, cogongrass is a C4 plant species. The C4 pathway allows grasses like cogongrass to photosynthesize more efficiently under

conditions of high light intensity, high temperatures, and limited moisture availability

(Groves, 1991).

Habitat

Cogongrass is widely distributed, it occurs at sites at sea level and at high •,

altitudes (Dickens and Buchanan, 1971). It is normally found in warm areas where the

annual rainfall is 75-500 cm (Bryson and Carter, 1993). Its habitat is diverse, that ranges

from the coarse sands of the desert dunes or along shorelines to the fine sands or sandy

loam soils of swamps and riverbanks. It thrives on fine sand to heavy clay and does well

on poor, low fertility soils. In Asia, stands of cogongrass are typically found on highly

leached soils with low pH, fertility, and organic matter (Sajise, 1980). Cogongrass is able

to establish monotypic stands in dry, poor, acidic soils. It survives dry soil conditions but

does not tolerate cool temperatures. Hubbard et al. (1944) reported that cogongrass

cannot survive temperatures of -8°C, however, Wilcut et al. (1988) documented rhizome

survival at -14°C. Cogongrass is adapted to full sun and it does not tolerate shade well

(Colvin et al., 1993). 15

Cogongrass Management

Chemical Control. Cogongrass is not a prevalent problem throughout the United States

and it is not a weed in most cropping areas, and thus there has been no impetus to develop herbicides specifically for its control (Shilling et al., 1997). As with other perennial weed species, successful management of cogongrass requires the deUvery of a lethal chemical dose to the underground tissues. At least 30 herbicides and hundreds of herbicide combinations have been tested and reported to control cogongrass (Bacon,

1986; Dickens and Buchanan, 1975; Southeast Asian Weed Information Center, 1988).

Imazapyr and glyphosate are effective herbicides to control cogongrass but a single application of either imazapyr and glyphosate does not successfully control cogongrass

(Akobundu, 1993; Dickens and Buchanan, 1975; Lee, 1985; Shilling et al, 1997; Shilling

et al., 1995; Sriyani, 1992; Tanner et al., 1992; Townson and Butler, 1990; Townson and

Price, 1987; Willard, 1988; Willard et al., 1997). Both herbicides are readily absorbed

and translocated throughout the plant (Dean et al., 1988). In most situations, long-term or complete control has not been achieved from a single application of any chemical herbicide. Multiple applications are necessary to inhibit the regrowth from the extensive rhizome system. Hence, the greatest levels of control were achieved with repeated or

sequential applications of imazapyr and glyphosate (Dean et al., 1988; Townson and

Butler, 1990; Willard et al., 1996). So far, an evenly split application of 4.5 kg/ha glyphosate and 2.2 kg/ha imazapyr is the most effective chemical management strategy for cogongrass (Willard et al., 1996). Control with this strategy is enhanced by discing the cogongrass before herbicide application to break apical dominance and stimulate growth of new leaves and by discing after herbicide application to incorporate herbicide

' ' • in the soil (Dozieretal., 1998). ,

Although multiple applications of glyphosate can provide substantial control of

cogongrass, its use in areas with desirable vegetation is limited by its non-selective nature, which creates the risk of injury to non-target plants. Application of imazapyr sterilizes the soil and prevents the growth of any vegetation for six months to a year, predisposing the treated area to erosion. The residual soil activity of imazapyr limits its use for cogongrass control (Colvin et al., 1994). Other problems associated with the use of chemical herbicides are increased annual costs, overdose application that results from ignorance of weed heterogeneity, and potential selection for herbicide resistant biotypes

' (Shilling et al., 1997). x :

Mechanical Control. Since cogongrass does not persist in frequently cultivated areas,

mechanical means of control have been used in certain situations, with varying levels of

success. Tillage reduces shoot growth and damages rhizomes. Deep and frequent tillage

is required to prevent rhizomes from producing new shoots for regrowth. Shallow tillage

only fragments rhizomes, causing short-term growth reduction and subsequent sfrong

shoot growth. The effectiveness of tillage also relies on the removal of foliage biomass.

Tillage done during the hot dry summer would result in higher rhizome mortality. The

use of tillage is limited to areas where erosion potential is low and where rhizomes will

not be transported beyond the infested area (Willard, 1988).

Mowing depletes rhizome reserves but the large rhizome reserves of cogongrass

allow it to produce new growth quickly after mowing, thus, sequential mowings are needed over two or more years to deplete the rhizome reserves eventually (Shilling et al.,

1995). Discing reduces shoot growth and damages rhizomes by chopping and desiccation. Discing alone has not been effective to control cogongrass although reduction in biomass has been reported (Gaffhey, 1996; Willard, 1988). Continuous discing may be effective but this type of mechanical control is not practical or feasible in many areas, such as natural sites that are not completely dominated by cogongrass.

Removal of old growth by burning or mowing causes rhizomes to reallocate starch

storage reserves to produce new shoots, which weakens the rhizomes. Burning or mowing also removes dead biomass and results in improved tillage (Dozier et al., 1998).

Cultural Control. In some countries, leguminous crops or tree hedgerows have been

used to control cogongrass which also increase nitrogen in the soil (Akobundu, 1993;

Cox and Johnson, 1993). Control with cover crops exploits the sensitivity of cogongrass,

a C4 plant, to shading. Alley cropping with leucaena (Leucaena leucocephala [Lam.] de

Wit) and mother of cocoa {Gliricidia sepium [Jacq.] Walp.) reduced rhizome dry matter

by >90% compared to natural fallow (Anoka et al., 1991). Other species that have been

used as cover crops include velvetbean {Mucuna pruriens [L.] DC), calop

{Calopogonium mucunoides Desv.), centro {Centrosema pubescens Benth.), tropical

kudzu (Pueraria phaseoloides [Roxb.] Benth.), stylo (Stylosanthes guyanensis), sunn

hemp {Crotolaria juncea L.), and jackbean {Canavalia ensiformis [L.]) (Manyong et al.,

1996; Versteeg and Koudokpon, 1990; Anonymous, 1995; Guritno et al., 1992). From

studies done in West Africa, control of cogongrass by planting velvetbean might be a

better alternative for farmers who cannot afford herbicides. 18

Integrated Management. There is no single treatment that effectively eliminates

infestations of cogongrass. Many authors have pointed out that reliance on a single

control measure will generally result in failure to manage cogongrass effectively (Byrd

and Bryson, 1999; Dozier et al., 1998; Shilling et al., 1995). The integration of chemical

and mechanical means of control has been considered a viable approach to control

cogongrass. Tillage and discing would reduce the rhizome system. Burning or mowing

would remove dead biomass and reduce the chance of herbicide interception by dead

tissues. Herbicide application to the flush of new plant tissue would result in more

efficacious control. A combination of discing followed by application of imazapyr,

glyphosate, or a mixture of both may provide greater than 90% control. A combination

of burning or discing and application of imazapyr was also effective (Colvin et al., 1994;

Dozier et al., 1998; Shilling et al., 1995).

Cogongrass is able to reinvade an ecological niche if it is not replaced by another plant species following control. Revegetation with desirable species after chemical

suppression is necessary to achieve long term control of cogongrass (Shilling et al.,

1997). Bermudagrass {Cynodon dactylon (L.) Pers.) and hairy indigo (Indigofera hirsuta

Harvey) in combination with glyphosate and imazapyr, respectively, grew vigorously and

suppressed cogongrass for up to 2 years after seeding. The success of this strategy depends on the tolerance of the desired species to the herbicide and soil type, among other factors (Gaffhey, 1996). The use of plant species to suppress re-establishment of cogongrass and displace it is limited by the lack of experience with the management of most desired species (Shilling et al., 1995). Biological Control. The use of biological control agents may be also essential to manage cogongrass, especially in low maintenance areas where the cost of using chemical herbicides and mechanical control measures cannot be justified. Cogongrass is

a good target weed for biological control because it forms monotypic stands and is primarily a problem in non-crop areas, hence the risk of non-target plant injury would be minimized.

The development and registration of several bioherbicides worldwide is indicative of the interest that the bioherbicide approach to control weeds has generated in recent years. In the 1980s, Devine®, composed of the Phytophthora palmivora [Butler] Butler, was registered for the control ofMorrenia odorata (Hook. & Am.) Lindl. in citrus in

Florida. A year later, another bioherbicide, CoUego , composed of the fungus

Colletotrichum gloeosporioides (Penz.) Penz. & Sacc. f sp. aeschynomene, was registered for the control Northern jointvetch {Aeschynomene virginica (L.) B.S.P.) in rice and soybeans in the United States (TeBeest and Templeton, 1985). Since the 1980s, several other bioherbicides have been developed, registered, and used on various weed species in several countries. Dr. Biosedge, composed of the rust fungus Puccinia caniculata (Schwein.) Lagerh. has been registered, but not currently used, for the control of yellow nutsedge {Cyperus esculentus L.) in the United States (Greaves and MacQueen,

1992). In Japan, Xanthomonas campestris Migula pv. poae has been registered as

Camperico , a bacterial bioherbicide for annual bluegrass {Poa annua L.) that occurs in turf and golf greens (Imaizumi et al., 1997). Another bioherbicide, CHONTROL, composed of the fimgus Chondrostereum purpureum (Pers. ex Fr.) is under development for North American registration and use for stump application in Canadian and U. S. 20

forests (Dumas et al., 1997; Prasad, 1994; Shamoun and Hintz, 1998). Colletotrichum

gloeosporioides f. sp. malvae (Mallet), a bioherbicide for round-leafed mallow (Malva pusilla Sm.) and small-flowered mallow (M parviflora L.), is under development (R.

Charudattan, personal comm.) and Altemaria destruens (Smolder®), a bioherbicide for

dodder (Cuscuta spp.), is undergoing EPA review for registration in the United States

(Environmental Protection Agency Office of Pesticide Programs, 2001).

Cylindrobasidium laeve (Pers: Fr.) Chamuris, wood-rotting basidiomycete, has been

registered in South Africa and is now being sold as Stumpout , a stump-treatment product to prevent regrowth of wattles {Acacia mearnsii de Wildman) and A. pycnantha

Benth., in the Cape Province (Morris et al., 1998).

The mycoherbicidal strategy, exemplified by the commercial use ofPhytophthora palmivora (De Vine) and Colletotrichum gloeosporioides f sp. aeschynomene (Collego), utilizes fiingal pathogens in a manner that is consistent with modem herbicide application technology and equipment. Applications of mycoherbicides are directed to the area of weed infestation and applied, using conventional application methods and tools, in volumes and dosages that suppress the population of target weeds within an allotted time before economic losses are incurred. The fimgal pathogen is usually applied as a formulated product and is not expected to provide weed control beyond the growing

season in which it is applied. Applications are required annually since the pathogen generally does not survive in sufficient numbers and does not multiply to provide the

inoculum needed to initiate a new epidemic on new weed infestations (TeBeest et al.,

1992; Charudattan, 1985; TeBeest and Templeton, 1985; Smith, 1986). Several pathogens have been found in association with cogongrass and some have

been evaluated for their potential as bioherbicides. Myrellina imperata Sankaran &

Sutton causes leaf spots and chlorosis, and Giberella imperatae C. Booth & Prior causes

dieback on cogongrass accessions from Papua New Guinea and Australia (Sankaran and

Sutton, 1991; Booth and Prior, 1984). Persley (1973) reported that the causal agent of

leaf scald of sugarcane, Xanthomonas albineans (Ashby) Dowson caused the appearance

of white and yellow lines along the leaf veins of cogongrass in Australia. In Malaysia,

two antrachnose-causing fiingi, {Colletotrichum caudatum (Sacc.) Peck and C.

graminicola (Ces.) G.W. Wils], Bipolaris maydis (Nisikado & Miyake) Shoemaker, and

the rust fungus, Puccinia rufipes Diet., were reported on cogongrass. The occurrence of

P. rufipes on cogongrass has also been reported in Thailand. The exploratory phase of

research on biological control in Malaysia has focused on the evaluation of

Colletotrichum caudatum, Ascochyta sp., Puccinia rufipes, C. graminicola, Didymaria

sp., and Dinesmasporium sp. as potential biological control agents for cogongrass. In

inoculations in the greenhouse with C. caudatum produced necrotic streaks along the leaf

edges, but plant death was not observed (Gaunter, 1996). In the United States, Chase et

al. (1996) evaluated several ftingal pathogens from diseased cogongrass collected from

various sites in Florida. Four of the six isolates tested were pathogenic to cogongrass and

caused 60-100% disease incidence, but weed mortality was not observed. Chaetomium

fusiforme Chivers was found to be the most promising, as it caused 100% disease

incidence in greenhouse trials. An isolate belonging to the Helmithosporium group

caused 85% disease incidence. The other fiangal pathogens isolated in Florida were

species of Curvularia, Fusarium, and Pestalotia (Chase et al., unpublished data). Other fungi that were reportedly found in association with cogongrass include

Aschochyta imperatae Punithalingam, Claviceps imperatae Tanda & Kawatani,

Monodisma fragilis Alcorn, Deightoniella africana Hughes, and Mycosphaerella imperatae Sawada (Van Dyke and Ravenell, 1985). Although a number of pathogens have been found to occur on cogongrass, there are no bioherbicides that are registered for the control of cogongrass at present.

The objectives of this study were to: 1) screen indigenous fungal pathogens of grasses for their biological control potential on cogongrass, 2) select a suitable pathogen to develop as bioherbicide, 3) develop materials and methods to mass-produce the inoculum, 4) determine the optimum conditions for inoculum storage to prolong the shelf-life of the bioherbicide, 5) determine a suitable formulation to increase biocontrol efficacy, 6) determine the host range and selectivity of the bioherbicide agent,

7) determine the effects of formulation and appUcation rate on the field performance of the bioherbicide, and 8) evaluate the suitability of biological control as a component of an integrated approach to manage cogongrass. CHAPTER 2 EVALUATION OF FUNGAL PATHOGENS AS BIOLOGICAL CONTROL AGENTS FOR COGONGRASS (Imperata cylindrica (L.) Beauv.)

* . Introduction

Integrated management of cogongrass is a promising, but a high-input approach, to displacement of this invasive weed by desirable grasses. The cost of

employing the integrated approach precludes its use in low-maintenance and natural areas where cogongrass frequently occurs. The use of biological control agents would

be ideal in these situations where high-input weed control will not work or is not

feasible (Dozier et al., 1998; Shilling et al., 1995).

Plant pathogens and insects have been considered as cost-effective alternative means to control weeds, such as cogongrass, that occur in natural ecosystems and low maintenance areas where the cost of chemical herbicides and the use of mechanical control methods are not justified or feasible. Chemical control methods, aside from being costly, also have the potential to cause injury to nontarget plants, soil residual activity, environmental pollution, and herbicide resistance. The use of plant pathogens to control weeds may be used in situations where a high degree of host

specificity is required and the use of chemicals is not suitable because of environmental contamination (Auld and Morin, 1995).

A number of insects and pathogens have been found in association with cogongrass in many countries of the world. Vayssiere (1957) reported several

23 24

polyphagous insects that attack cogongrass as well as other cultivated cereals. Three

species of the North American skipper butterfly (Lepidoptera: Hesperiidae),

Ancyloxypha numitor (Fabricus), Atalopedes campestris (Boisduval), and Hylephila phyleus Drury, have been reported to feed on cogongrass (Bryson, 1985; Bryson,

1987). These three species were found to be unsuitable agents for biological control

because their larvae also feed on several grassy crop species that include

bermudagrass {Cynodon dactylon (L.) Pers.), com {Zea mays L.), and St. Augustine

grass (Stenotaphrum secundatum (Walt.) Ktze.) (Bryson and Carter, 1993).

Cogongrass is also a host of the skipper butterfly (Pelopidas mathias oberthueri

Evans.) (Lepidoptera: Hesperiidae), a pest of rice in Japan (Masuzawa et al., 1983).

Several pathogens have been reported on cogongrass, including Myrellina

imperatae Sankaran & Sutton, Giberella imperatae C. Booth & Prior (Sankaran and

Sutton, 1992; Booth and Prior, 1984), Xanthomonas albilineans (Ashby) Dowson

(Persley, 1973), Puccinia rufipes Diet. (Chadrasrikul, 1962), Claviceps imperatae

Tanda & Kawatani, Monodisma fragilis Alcorn, Deightoniella africana Hughes,

Mycosphaerella imperatae Sawada, Bipolaris maydis (Y. Nisik.) Shoemaker,

Colletotrichum caudatum (Sacc.) Peck, C. graminicola (Ces.) G. W. Wilson,

Aschochyta sp., Didymaria sp., Dinesmasporium sp. (Caunter, 1996), Chaetomium

fusiforme Chivers, and species of Helminthosporium, Curvularia, Fusarium, and

Pestalotia (Chase et al., 1995, unpublished data). Cogongrass is an alternate host of

Rotylenchus reniformis (Linford & Oliveira) (Anonymous, 1978), it is also a host of

the rice-tungro virus (RTV) and associated viruses (International Rice Research

Institute, 1988). 25

The biological control potential of C. caudatum, Ascochyta sp., P. rufipes, C.

graminicola, Didymaria sp., and Dinemasporium sp. were investigated in Malaysia

(Gaunter, 1996). Inoculations in the greenhouse with C. caudatum reportedly caused necrotic streaks along the leaf edges of cogongrass but plant death was not observed

(Gaunter, 1996). In the United States, Ghase et al. (1996) evaluated several fungal isolates as potential biological control agents for cogongrass. Four of the six isolates tested were pathogenic to cogongrass and caused 60-100% disease incidence but weed mortality was not observed.

Presently, there are no biological control agents registered for use in the classical biocontrol way or the bioherbicide approach to control cogongrass. This

situation may be due in part to the lack of a pathogen that is sufficiently aggressive to cause significant damage on cogongrass. The objectives of this study were: 1) to

screen indigenous fungal pathogens of grasses for their potential as biological control

agents of cogongrass, and 2) to select a suitable pathogen for development as a

bioherbicide.

Materials and Methods

Gollection and Isolation of Fungal Pathogens

Samples of diseased plants were collected from stands of cogongrass in

Gainesville and Waldo, Florida in 1996-1997. Diseased leaves were cut into 4 mm^

pieces, surface sterilized with 0.5% sodium hypochlorite solution for three minutes,

rinsed twice with sterile water, blot-dried on clean paper towels, and placed on potato

dextrose agar (PDA) in petri plates (100 x 15 mm). Agar plates that contained the 26

leaf pieces (5 pieces/plate) were incubated in the dark for 5-7 days at 27°C. Fungal growth from each of the diseased leaf pieces was aseptically transferred to fresh PDA plates and allowed to grow. Hyphal tips from the PDA cultures were transferred to

PDA slants and incubated for 7-10 days at 27°C to obtain pure cultures of the

isolates. Slants of pure cultures were stored in a 10°C incubator.

Pure cultures of other fungal pathogens were obtained from the culture

collection of Dr. Charudattan (Plant Pathology Department, University of Florida,

Gainesville, FL). These pathogens had been isolated from diseased grasses collected

in Florida and have been evaluated for their pathogenicity on several species of

weedy grasses.

Production of Inoculum

To produce inoculum, mycelial blocks from the slant cultures were transferred

to fresh PDA plates and allowed to grow for 5-7 days. Mycelial blocks from these

plate cultures were used to inoculate PDA or V-8 juice agar (100 ml V-8 juice/1 of

media) in several 100 x 15 mm petri plates. The plates with V-8 juice agar and PDA

were incubated in the dark at 27°C for 14 days. After the plates were fully covered

with mycelial growth, the cultures were checked for spores by making slide mounts

of scraped mycelia. Inoculum from sporulating cultures was harvested by adding 10

ml of 1% aqueous gelatin (Knox® unflavored gelatin; Nabisco Inc., East Hanover,

NJ) solution on each agar plate culture and the surface was gently scraped with a

rubber policeman. The gelatin solution was used to improve adhesion of the spores to

the leaves upon inoculation. The suspension of spores and mycelial fragments was 27

filtered through two layers of clean cheesecloth to remove mycelial debris. The spore concentration was determined with the aid of a hemacytometer.

Cultures that did not produce spores on V-8 juice agar or PDA were grown on

V-8 juice in flasks that were incubated on a shaker at room temperature for 7-14 days.

Fresh mycelium was harvested from the flasks and placed on a clean weighing boat.

The pre-weighed mycelium was blended for 10 seconds with the appropriate amount of sterile water in a Waring commercial blendor (Waring Products Division, New

Hartford, CT). Depending on the amount of mycelium available, the ratio of mycelium to water ranged from 0.1-0.5 g mycelium/ml of sterile water. Metamucil®

(0.5%; psyllium mucilloid; Procter & Gamble, Cincinnati, OH) was added to the mycelial suspension as a humectant to prevent desiccation after application on the leaves.

Propagation of Test Plants in the Greenhouse

- Cogongrass rhizomes were collected from a cogongrass stand near Lake

Alice, University of Florida Campus, Gainesville and brought to the greenhouse. The

rhizomes were cut into 4-cm segments and planted in plastic trays (13 x 17 x 56 cm)

that contained a commercial potting medium (Metromix 300; Scott' s-Sierra

Horticultural Products Co., Marysville, OH). The trays were fertilized with a 14-14-

14 fertilizer (Osmocote; Scott' s-Sierra Horticultural Products Co., Marysville, OH).

After 2-4 wks, rhizomes with healthy shoots were transplanted into clay pots (1 1.5-

cm diameter; 1 1 .0-cm height). 28

Inoculation

Inoculations were carried out in the greenhouse four to five weeks after the rhizomes were sown. Spore suspensions were applied until run-off with hand-pumped

sprayers. Mycelial inoculum was applied to cogongrass leaves with sterile

paintbrushes. All inoculated plants and nontreated control plants were incubated in a

dew chamber for 24 h (27 + TC, 100% RH) and then transferred to the greenhouse

bench for observations. The experiments consisted of three replications per

treatment.

Pathogenicity Testing

Inoculated plants were observed for symptom appearance within 7 days after

inoculation (DAI). Disease symptoms produced by each of the isolates were

characterized and diseased tissues were plated on PDA in petri plates for isolation of

the causal organism. Fungi that were isolated from diseased tissues were compared

with the original isolates used in the inoculation for cultural characteristics and

morphology. The Koch's postulates were fulfilled.

Identiflcation of Isolates

The genera of unknown isolates collected from the field were identified on the

basis of their morphological characteristics. Dr. James W. Kimbrough (Plant

Pathology Department, University of Florida, Gainesville, FL) identified the

promising biocontrol isolates to the species level. Non-sporulating isolates were

excluded from further evaluation due to their inability to produce spores readily on 29

common laboratory media and because they could not be identified based on mycelia alone.

Efficacy of Promising Biocontrol Isolates Applied Singly or as a Mixture

In initial tests, some of the fungal isolates caused mere speckling or lesions.

These isolates were not considered sufficiently aggressive and were excluded from further testing. The isolates that caused coalescent lesions and blighting, Bipolaris

sacchari and Drechslera gigantea, were tested further. These two isolates were

evaluated individually and combined for their efficacy in the greenhouse.

The spore concentrations of B. sacchari, D. gigantea, and their mixture were 3

X 10^, 5 X 10^, and 1.5 x 10^ spores/ml for the first, second, and third trials,

respectively. The spores were suspended in sterile water with 1% unflavored gelatin

as an adjuvant. Inoculated and control plants (treated with 1% gelatin only) were

incubated in a dew chamber for 24 h (27 ± 1° C, 100% RH) and then transferred to

the greenhouse for further observation. The bioherbicidal efficacy of the isolates was

measured in terms of percent disease severity (DS). The DS level was assessed with

the aid of a pictorial key for southern com leaf blight (James, 1971), with 50% as the

maximum level of disease. Assessments were done at 7 and 21 days after inoculation

(DAI). The experiment had a completely randomized design with three replications

per treatment. 30

Statistical Analysis

Percentage data were transformed using an arcsine square root transformation.

Analysis of variance (ANOVA) was performed using the General Linear Models

Procedure (SAS Institute, Gary, N.G.)- Means were compared using the Duncan's

Multiple Range test at P=0.05.

Results

GoUection, Isolation, and Determination of Pathogenicity of Fungal Pathogens

Diseased cogongrass samples collected from Gainesville and Waldo, Florida yielded seven fungal isolates. Three of these isolates did not produce spores when grown on PDA or V-8 juice agar. Gultures of these isolates on PDA were characterized by fluffy, whitish-gray mycelial growth. The four other isolates from cogongrass produced spores on PDA. The sporulating cultures were identified as

Chaetomiumfusiforme (isolated by G. Ghase; provided by Dr. Gharudattan),

Bipolaris sacchari, and two Curvularia species. The other isolates, namely,

Drechslera gigantea (from Digitaria sanguinalis [L.] Scop.), Exserohilum

longirostratum (from Dactolectenium aegyptium [L.] Willd.), and E. rostratum (from

Sorghum halepense [L.] Pers.) were provided by Dr. S. Ghandramohan (Plant

Pathology Department, University of Florida, Gainesville, FL). Another

nonsporulating isolate was obtained from a diseased specimen of Echinochloa species

collected during a field trip to Dade Gounty, Florida in 1996.

All 1 1 fungal isolates were pathogenic on cogongrass in greenhouse tests.

The pathogenicity of these isolates was confirmed by fulfilling Koch's postulates. 31

Disease symptoms ranged from minute specks to leaf blight (Table 2-1). Of the seven sporulating isolates tested, Chaetomium fusiforme and the two species of

Curvularia only caused minute specks on the leaves. Exserohilum rostratum and E. longirostratum caused discrete and coalescent leaf lesions. Drechslera gigantea and

Bipolaris sacchari caused discrete and coalescent lesions and leaf blight (Fig. 2-1).

Bipolaris sacchari and D. gigantea were chosen for further evaluation based on the disease symptoms they caused on cogongrass in these initial inoculations in the greenhouse.

Biocontrol Efficacy of Bipolaris sacchari and Drechslera gigantea Applied Singly or as a Mixture

Since B. sacchari and D. gigantea caused substantially higher levels of DS than the other isolates, they were studied further. At the early stages of the disease they caused, discrete lesions that were brown to dark reddish brown in color, elongate or somewhat spindle-shaped with white or pale centers and brown tapering ends. The lesions ranged from 1.0 to 2.0 mm in length and did not exceed 0.5 mm in diameter.

As the disease progressed, the areas around some of the discrete lesions turned yellow. Lesions that were very close together appeared as one large, irregularly shaped necrotic lesion due to the yellow borders that eventually turn brown.

The discrete lesions did not have a dramatic increase in size (the diameter of some lesions increased from 0.5 to 0.8 mm, and lengths by 0.2 to 0.5 mm) but the yellowing of the borders became widespread and caused large areas to turn necrotic.

Tips and edges of infected younger leaves turned dark green to brown and eventually dried up, giving the leaf a folded appearance. Older leaves exhibited more severe 32

disease symptoms: large necrotic areas with yellow borders and subsequent blighting of most of the leaf blade, beginning at the tips and edges of leaves, were observed within 72 h after inoculation.

No secondary disease spread was observed in the greenhouse. Both isolates caused plant death but high levels of disease did not always translate into high mortality of weed. The infection caused by the isolates was localized and limited to the sprayed foliage. Newly emergent leaves of inoculated plants were disease-free and the healthy regrowth decreased the level of DS after a few weeks.

The repeated trials were analyzed separately due to a significant treatment by

trial interaction effect (Table 2-2). In the first trial, treatment had a significant effect at 7 and 21 DAI. B. sacchari, D. gigantea, and the mixture caused 47.2%, 43.5%, and 48.8% disease severity at 7 DAI, respectively. Both B. sacchari and the mixture caused the higher DS than D. gigantea. At 21 DAI, the DS caused by B. sacchari, D. gigantea, and the mixture dropped to 42.6%, 35%, and 47.7%. The lowest DS was observed in plants treated with D. gigantea (Fig. 2-2 A).

In the second trial, there was no significant treatment effect at 7 and 21 DAI.

B. sacchari, D. gigantea, and the mixture caused similar levels of disease. The DS ranged from 43.8% to 47.5% at 7 DAI and from 37.6% to 42.1% at 21 DAI (Fig. 2-2

B). Similar results were obtained from the third trial. DS levels caused by all three treatments were similar at 7 DAI (DS levels ranged from 42.4% to 45.8%) '

33

Table 2-1. Disease symptoms caused by fungal pathogens tested on cogongrass {Imperata cylindrica [L.] Beauv.)

Inoculum Pathogen Isolated From Collected Concentration Cogongrass From Tested Reaction

^ Bipolaris sacchari Cogongrass Gainesville, 10^ spores/ml Lesions and FL blight

Chaetomium Cogongrass Florida 10^ spores/ml Specks ^ fusiforme

Curvularia sp. Cogongrass Gainesville, 10^ spores/ml Few specks Isolate #13*' FL

Curvularia sp. Cogongrass Gainesville, 10* spores/ml Numerous Isolate #15 FL specks

Drechslera Crabgrass Leesburg, FL 10^ spores/ml Lesions and ^ gigantea blight

Exserohilum Crowfootgrass Belleglade, 10^ spores/ml Lesions ^ longirostratum FL

Exserohilum Johnsongrass Gainesville, 10^ spores/ml Lesions ^ rostratum FL

Isolate # 3 Cogongrass Gainesville, 0.5 g mycelia/ml Lesions and FL blight

o _ 1 iSOldlC TT Cogongrass Gainesville, 0. 1 g mycelia/ml Specks FL " Isolate # 8 Echinochloa Dade County, 0.5 g mycelia/ml Lesions sp. FL Isolate # 10 Cogongrass Gainesville, 0.4 mycelia/ml Lesions and FL blight

" Isolates that were further evaluated for biocontrol efficacy on cogongrass. Isolates that were not considered further as biocontrol agents because of the mild symptoms they caused. Nonsporulating isolates that were excluded from further consideration because they could not be easily identified. 34

Fig. 2-1. Disease symptoms caused by Bipolaris sacchari (top) and Drechslera gigantea (bottom) on cogongrass. The symptoms were observed within 7 days after inoculation (DAI). 35

Table 2-2. Analysis of variance for disease severity of cogongrass inoculated with Bipolaris sacchari, Drechslera gigantea, and their mixture Source DF SS MS F Value Pr>F

Treatment 2 0.01702923 0.00851462 4.57 0.0158

Trial 2 0.00753544 0.00376772 2.02 0.1446

Replication 2 0.00263343 0.00131671 0.71 0.4987

Treatment x trial 4 0.02304425 0.00576106 3.09 0.0252

DF = degrees of freedom. SS = sum of squares. MS = means square error. 36

B. sacchari D. gigantea Mixture Treatment

B

B. sacchari D. gigantea Mixture Treatment

B. sacchari D. gigantea Mixture Treatment

Fig. 2-2. Disease severity caused by Bipolaris sacchari and Drechslera gigantea at 7 and 21 days after inoculation (DAI). The pathogens were applied individually or combined. Within each DAI, bars with similar letters are not significantly different at

P=0.05, as determined by Duncan's Multiple Range test. Results from three trials are shown (A, B, and C). 37

and at 21 DAI (Fig. 2-2 C). The DS ranged from 42.4% to 45.8% at 7 DAI and from

35.1% to 42.4% at 21 DAI.

Bipolaris sacchari and D. gigantea caused high levels of disease but did not consistently cause plant death. At 21 DAI, the percentage weed mortality (% WM) caused by B. sacchari was 33.3%, 26.7%, and 40%, for the first, second, and third

trials, respectively. Application of D. gigantea caused 26.7% and 13.3% weed

mortality in the second and the third trials but plant death was not observed in the first

trial. The mixture of the two pathogens caused plant death during the first trial

(66.7%) and the second trial (40%), but there was no plant mortality for the third trial.

There was a decrease in disease severity at 21 days after inoculation due to the emergence of new healthy leaves. Aside from the regrowth, new shoots also emerged from under the soil after 21 days after inoculation. The application of the fungal pathogens did not adversely affect the underground reproductive structures of cogongrass.

Discussion

The potential of the fungal pathogens Bipolaris sacchari (E.J.Butler) Shoem. and Drechslera gigantea (Heald & F. A. Wolf) Ito as biological control agents of cogongrass (Imperata cylindrica [L.] Beauv.) was confirmed in repeated greenhouse trials. Four- to 5-week-old cogongrass plants inoculated with either isolate exhibited lesions within 24- 48 h after inoculation and incubation in a dew chamber. The disease symptoms caused by the isolates were similar in appearance (Fig. 2.1). 38

Bipolaris sacchari (formerly Helminthosporium sacchari [Van Breda de

Haan] Butler) has been known as the causal organism of eyespot and seedling blight

of sugarcane (Saccharum officinarum L). This fungus produces a compound that is selectively toxic to susceptible sugarcane clones (i.e., host specific) (Scheffer and

Livingston, 1980; Steiner and Strobel, 1971). The Bipolaris sacchari (HS) toxin causes the formation of runner lesions and electrolyte leakage from sugarcane clones

susceptible to B. sacchari (Livingston and Scheffer, 1984). Zhang et al. (1996) showed that a toxin produced by B. sacchari in culture was biologically active on

Echinochloa species. When placed in cell-free filtrates of B. sacchari for 24 h, intact seedlings exhibited chlorosis and wilting, symptoms similar to plants that were inoculated with spore suspensions of B. sacchari.

Bipolaris sacchari is also reported to cause leaf diseases of lemon grass

{Cymbopogon citratus [D. C. ex Nees] Stapf) and napiergrass (Pennisetum pupureum

Schumach). Cross inoculations of B. sacchari from lemon grass and napiergrass to sugarcane only produced minor infections, which was interpreted as existence of biotypes (Cook, 1981). In India, B. sacchari was isolated from eyespots on Imperata

arundinacea Cyr. (Misra, 1973). Bipolaris sacchari is indigenous to Florida where it has been reported on pampasgrass (Cortaderia selloana [Schult. & Schult.f ] Asch. &

Graebn.), lemongrass {Cymbopogon citratus [DC] Stapf), ravennagrass {Erianthus ravennae Beauv.), guineagrass (Panicum spp.), napiergrass (Pennisetum pupureum

Schumach.), and sugarcane (Saccharum officinarum L.) (Alfieri et al., 1994).

Bipolaris sacchari has been isolated from leaf spots on weedy P. maximum Jacq. 39

(narrow leaf Panicum) in citrus groves in Florida. The same isolate had little effect

on another Panicum variety, P. maximum var. trichoglume (Sonoda et al., 1992).

Bipolaris sacchari is one of four pathogens that have been evaluated for their biocontrol potential on Echinochloa species occurring in rice. In inoculations done in the greenhouse, B. sacchari caused 100% disease severity on E. colona (L.) Link and

E. glabrescens Munro ex Hooker after 2 and 8 DAI, with 24-h dew exposure (Zhang et al., 1996). Under controlled environments, other species of Bipolaris have shown potential as biocontrol agents of other serious grass weeds. The application of B. setariae (Saw.) Shoemaker on goosegrass (Eleusine indica [L.] Gaertn.) resulted in flecks that expanded and coalesced to cover large areas of the leaf. Lesions on the stem caused girdling and plant death. High levels of disease were achieved when

inoculated plants were provided with 48 hours of dew at 24°C (Figliola et al., 1988).

In another study, B. setariae caused systemic infection and 90% plant mortality of 17- day-old seedlings of broad-leaf signal grass (Brachiaria platyphylla [Griseb.] Nash) after 8 h of dew exposure in the greenhouse and laboratory (Ravenell and Van Dyke,

1986). Two other species of Bipolaris, B. halepense and B. sorghicola (Lefebvre &

Sherwin) Alcorn, were also considered as potential biocontrol agents of johnsongrass

(Sorghum halepense [L.] Pers.) (Chiang et al., 1989a: Chiang et al., 1989b; Winder and Van Dyke, 1987; Winder and Van Dyke, 1990).

Drechslera gigantea (formerly Helminthosporium gigantea [Heald. & Wolf]) is the causal organism of the zonate eyespot disease on a number of grass genera including Agropyron, Digitaria, Cynodon, Echinochloa, Panicum, and Pennisetum

(Drechsler, 1928; HoUiday, 1980). Zonate eyespot disease also occurs on wheat, rye. 40

and barley in the coastal areas near the Gulf of Mexico in the USA, Mexico, and

Central America (Zillinsky, 1983). Drechlsera gigantea also infects coconut and

Saccharum spp. and is the causal organism of the eyespot disease on banana in

Jamaica (Meredith, 1963). This pathogen is indigenous to Florida, and it causes zonate eyespot on bermudagrass and goosegrass, and leaf spot on crabgrass and

geranium (Pelargonium x hortorum L.H. Bailey) (Alfieri et al., 1994).

Drechlsera gigantea is known to produce numerous phytotoxins that belong to the eremophilane family. Two of these phytotoxins, petasol and gigantenone,

cause necrotic lesions on dicotyledonous species (Bunkers et al., 1990; Kenfield et

al., 1989) and induce the formation of green islands or localized areas of chlorophyll

retention in monocotyledonous species (Bunkers et al., 1991; Kenfield et al., 1989).

In the study by Bunkers et al. (1991), production of eremophilane by D. gigantea increased with the addition of plant extract of quackgrass (Agropyron repens [L.]

Beauv.), that was interpreted as the host induction of toxin production by the pathogen.

Toxin production, which occurs in several fungal genera such as Bipolaris

(teleomorph Cochliobolus), may be related to the strategy of pathogenicity of these organisms. Some species and races of Helminthosporium are primarily saprophytes that can be opportunistic pathogens. They have low virulence and can only infect senescent tissue or infect through wounds. Other species that belonged to the erstwhile genus Helminthosporium have been transferred to Bipolaris, Curvularia,

Drechslera, and Exserohilum. Contrarily, some other species and races of

Helminthosporium are primarily pathogens but can also exist as saprophytes under 41

special conditions and in culture. Such species are usually host-selective and virulent on their selected hosts. The ability to produce toxic compounds involved in disease development allows some species in the Helminthosporium complex to become more specialized on certain hosts and more adapted to the pathogenic lifestyle as compared to other organisms (Scheffer, 1991).

A number of Bipolaris species that have caused serious diseases on cultivated graminicolous species are toxin producers. Bipolaris victoriae F. Meehan & Murphy, causal organism of Victoria blight, produces victorin or HV toxin (Scheffer, 1976).

This toxin is extremely active against oats with the Vb gene but does not affect

resistant oat varieties or other species (Macko et al., 1989; Scheffer, 1976). Bipolaris

carbonum Ullstrup race 1, causal organism of com leaf spot, produces the HC toxin

(Scheffer, 1976). The pathogenicity and selectivity of C. carbonum depends on the excretion of the HC toxin, which is highly pathogenic to com that has the hm gene.

Bipolaris (Helminthosporium) maydis (Nas. & Miy.) Shoem. race T produces the

HmT toxin (Hooker at al., 1970; Yoder, 1976) and causes southern com leaf blight, a devastating disease of com lines with Texas male sterile (Tms) cytoplasm. The basis of virulence and selectivity by race T is the excretion of the HmT toxin (Scheffer,

1991). Bipolaris sacchari appeared suddenly and became a serious pathogen in

Hawaii, Puerto Rico, Java, and other parts of the world where sugarcane clones were planted (Scheffer, 1991). Like other members in the Helminthosporium complex, the pathogenicity and selectivity of B. sacchari isolate from sugarcane depend on the production and release of a host selective toxin (Scheffer and Livingston, 1980). 42

Chandramohan (1999) evaluated D. gigantea for its potential as a biocontrol agent for grass weeds in citrus groves. Drechslera gigantea controlled several grassy weeds when applied singly or in a mixture with two other grass pathogens,

Exserohilum longirostratum (Subr.) Sivan. and E. rostratum (Drechs.) Leon. &

Suggs. Grassy weeds that were found to be highly susceptible (82.5-100% severity) to each of the pathogens and their mixture include large crabgrass, crowfootgrass

{Dactyloctenium aegyptium [L.] Willd.), johnsongrass, guineagrass (Panicum maximum Jacq.), southern sandbur {Cenchrus echinatus L.), Texas panicum

(Panicum texanum Buckley), yellow foxtail (Setaria pumila [Poir.] Roem. & Schult.), green foxtail (S. viridis [L.] P. Beauv.), vaseygrass (Paspalum urvillei Steud.), natalgrass (Rhynchelytrum repens [Willd.] Hubbard), napiergrass (Pennisetum pupureum Schumach.), bermudagrass (Cynodon dactylon [L.] Pers.), torpedograss

{Panicum repens L.), and one variety of bahiagrass {Paspalum notatum Flugge). The

pathogen mixture was either superior or comparable to the individual pathogens in its biocontrol efficacy (Chandramohan, 1999). The use of a pathogen mixture did not necessarily result in significant increase in disease severity on most of the grass weeds.

Significant increase in bioherbicidal efficacy was not observed by Chiang et al. (1989b) when they inoculated johnsongrass seedlings with mixtures of E. turcicum

(Pass.) K. J. Leonard & E. G. Suggs and Colletotrichum graminicola (Ces.) G. W,

Wils and E. turcicum and Gloecercospora sorghi Bain & Edgerton ex Deighton.

Plzints treated with E. turcicum alone exhibited similar levels of disease as those treated with the pathogen mixtures. In the study reported here, control of cogongrass 43

was not necessarily enhanced by the application of the B. sacchari and D. gigantea mixture.

The two pathogens that have been selected as potential biocontrol agents for cogongrass are known to attack several other economically important, weedy, and nonweedy graminaceous species. Their ability to produce toxins may render them virulent control agents for cogongrass, as well as other non-target grass species.

However, since cogongrass often occurs in monotypic stands, the use of pathogens

that are not highly specific to cogongrass is not expected to pose the danger of injury to nontarget plants if directed, inundative inoculations are employed.

Although the isolates caused severe foliar disease, plant mortality was not always observed. Some inoculated plants produced healthy regrowth a few weeks after inoculation and new shoots emerged from under the soil. Thus, the ability of the rhizomes to generate new plants was not adversely affected by the inoculation with the pathogens. The level of damage that these isolates can cause in the field may be the same as that in the greenhouse test or lower, because field conditions may not be as favorable. However, biological efficacy of potential agents may be improved with appropriate formulation and timing of application. In cases where complete control of the target weed cannot be achieved by biological control alone, other control measures can be used in conjunction with the biological control agents. CHAPTER 3

MASS PRODUCTION OF INOCULUM OF Bipolaris sacchari and Drechslera gigantea AND DETERMINATION OF OPTIMUM CONDITIONS TO MAINTAIN SPORE VIABILITY DURING STORAGE

Introduction

The bioherbicide approach to weed control involves the application of massive amounts of infective inoculum to create an epidemic within the target weed

population in a short period of time. To be able to accomplish this, it is necessary to mass-produce viable, infective, and stable propagules of the pathogen through

inexpensive, efficient, and reproducible methods. This is a major requirement in the development of a bioherbicide (Churchill, 1982; Auld and Morin, 1995). Mass- produced propagules should also be amenable to long-term storage as dry preparations. Tolerance to desiccation and stability as a dry preparation are an

absolute requirement for commercial use of bioherbicides (Jackson et al., 1996;

Jackson, 1997). Zomer et al. (1993) claimed an overall lack of commercial success

of microbial biocontrol agents and attributed it to the difficulties in production and stabilization of these biocontrol agents, in addition to the lack of consistent efficacy in the field.

In general, there are three methods to produce inoculum: 1) the use of living host plants, 2) solid-substrate fermentation and, 3) liquid-culture fermentation. The use of living hosts for inoculum production is usually employed for the rust fungi,

44 45

which cannot be cultured on artificial media. High production costs, difficulties in propagule collection, and quality control of propagule characterize this method and

make it an unsuitable commercial option (Jackson, 1997).

The submerged liquid-fermentation system is the most economical method for

mass production of inoculum and is a readily available technology in most industrial countries (Churchill, 1982; Stowell, 1991). Submerged fermentation involves the use of a nutritional environment that can be maintained and monitored. The liquid medium simplifies the methods of production and processing and aids in the development of optimized nutritional conditions for production. Controlled nutritional and environmental conditions, process scale-up capabilities, quality assurance issues, and ease of product recovery contribute to lower costs when fungal propagules are produced with the liquid culture methods (Jackson, 1997). Submerged liquid-fermentation has been used for the commercial production of some of the registered or soon-to-be registered bioherbicides, such as Colletotrichum

gloeosporioides f. sp. aeschynomene (CoUego), Phytophthora palmivora (DeVine),

and C. gloesporioides f. sp. malvae (Mallet).

Submerged fermentation systems, although well developed for the production of bacterial cells or secondary metabolites such as antibiotics, amino acids, and ethanol, are not necessarily suited for mass production of fungal infective propagules

(Auld and Morin, 1995). Because of this, alternative methods to produce large amounts of inoculum with solid substrates have been developed for fungi that do not

readily produce conidia in liquid media. The use of solid-substrate fermentation is often the first method tested for mass production of inoculum since most fungi readily sporulate on solid substrates. Solid-substrate fermentation is easily accomplished

under laboratory conditions and the inocula produced tend to be more tolerant to

desiccation and more stable as dry preparation compared to conidia produced through

submerged liquid-fermentation (Jackson, 1997; Bartlett and Jaronski, 1988; Silman et

al., 1993). However, lower yields and longer fermentation times, which can translate

to higher production costs, render this system costly for commercial use (Jackson,

1997).

Methods for solid-substrate fermentation with agar media or grains have been

devised to produce the inoculum required to conduct greenhouse and field studies

with fungal pathogens. Conidia of Bipolaris zeicola (G. L. Stout) Shoemaker and B.

maydis (Nisikado & Miyake) Shoemaker were produced with finely ground, dry,

baby lima beans and crude agar in plates. The inoculated plates were incubated for

14 days, dried at 32-35°C to a thin film, and the spores were vacuumed using a

cyclone spore collector. This method produced 82 mg spores per plate, and each

plate contained 20 ml of the agar medium (Foudin and Calvert, 1987). Conidia of

three bioherbicidal fungi, Drechslera gigantea, Exserohilum longirostratum, and E.

rostratum, were mass-produced using V-8 juice agar on aluminum trays inoculated

with mycelial inoculum. This method produced spore suspensions (250 ml/tray) that

contained 2.59 x 10^ 5.47 x 10^ and 1.89 x 10^ spores per ml of Drechslera gigantea, Exserohilum longirostratum, and E. rostratum, respectively, 24- 48 h after the media were inoculated (Chandramohan, 1999).

Solid-substrate cultivation with grains instead of agar media has been utilized

in the mass production of Coniothyrium minitans Campbell, a fungus that parasitizes 47

sclerotia and mycelium of Sclerotinia sclerotiorum (Lib.) de Bary, a serious pathogen

of several economically important crops (Whipps and Gerlagh, 1992). Coniothyrium

minitans does not produce abundant conidia under submerged fermentation in stirred-

tank reactors but readily produced numerous spores on oats, perlite, or wheat bran

(Weber et al., 1999). Whole barley grains have been used to produce inoculum of

Beauveria brongniartii (Sacc.) Fetch., an entomopathogenic fungus that specifically

attacks cockhafers (Melolontha melolontha L.). Cultures of this fungus on barley

grains yielded 1 x 10^ to 2 x 10^ conidia per gram of grain in 24-30 days (Aregger,

1992). Inoculum of two other insect pathogens, B. bassiana (Balsamo) Vuillemin

and Metarrhizium anisopliae (Metschnikoff) Sorokin, have been mass-produced on

wheat bran (McCoy and Carver, 1941; Fogal et al., 1986). Daoust and Roberts

(1983) also produced conidia of M. anisopliae using three different types of rice.

Silman et al. (1991) compared four methods of inoculum production for

Colletotrichum truncatum (Schwein) Andrus & Moore, a potential bioherbicide for

hemp sesbania (Sesbania exaltata [Raf.] Rydb. ex A. W. Hill), by: 1) dialysis

membranes, 2) liquid media in shake flasks, 3) solid particles with humidity control,

and, 4) solid particles without humidity control. The spore yield for all four

substrates was lO' spores/ml of medium (Silman et al., 1991). While the quantity of

spores produced did not differ when solid substrates or liquid media were used,

spores that were dried after harvest from the solid substrate retained efficacy during

storage better than the spores that were dried after recovery from liquid cultures

(Silman et al., 1993). The choice of growth medium is important because of its influence on the quantity, type, stability, viability, and pathogenicity of the 48

bioherbicidal propagules (Bannon et al., 1990; Jackson and Bothast, 1990; Jackson

and Schisler, 1992; Morin et al., 1990; Schisler et al., 1991).

Spore stability, viability, and pathogenicity are also influenced by the storage

methods employed. The effects of temperature and humidity during storage on spore

viability have been studied for some fungal pathogens. Dried spores of C. truncatum

produced on liquid and vermiculite retained viability when stored at 4°C while those

produced on perlite-commeal agar and stored at 15°C retained the highest viability

(Silman et al., 1993). Conidia of Altemaria macrospora A. Zimmerm., a potential

biocontrol agent for spurred anoda (Anoda cristata [L.] Schltdl.), maintained over

90% viability for 12 months when stored in closed containers and refrigerated at 4°C

or frozen at -10°C (Walker, 1980). Generally, fungal spores stored in a dark, dry, and cool environment remain viable the longest. Storage of spores under optimal

conditions is therefore a key to prolong the shelf life of a bioherbicide.

The objectives of this study were: 1) to evaluate the suitability of a solid-

substrate culture method for the mass-production of B. sacchari and D. gigantea conidia, 2) to determine the effect of short-term storage methods on conidial viability,

3) to determine the effect of drying on the conidial viability, and 4) to determine the optimum relative humidity and temperature that would maintain the viability of dry conidia during storage. Materials and Methods

Evaluation of Grains as Substrates for Mass Production of Spores

Grains of rye {Secale cereale L.), oat (Avena sativa L.), and sorghum

{Sorghum bicolor [L.] Moench) were rinsed with running water to remove dirt. The grains were drained and weighed separately and 25 g of grains and 15 ml water were dispensed in glass bottles (5-cm diameter; 19.5-cm height). The bottled grains were autoclaved for 40 minutes at 121°C, at 15 psi (775.6 mm Hg), for two consecutive days. After cooling, the grains were inoculated with 5 ml (first trial) or 10 ml (second

trial) of blended mycelium of B. sacchari or D. gigantea with 500 |il of an antibiotic solution containing streptomycin sulfate (3.7 mg/ml) and chloramphenicol (2.5 mg/ml). The mycelium was harvested from 7-day-old V-8 juice cultures and blended for 10 seconds using a Waring commercial blendor (Waring Products Division, New

Hartford, CT). Media in bottles were inoculated and incubated at 27°C for 14 days, with alternating light and dark periods (12 h each). There were three replications per treatment.

On day 15, the spores were harvested by the addition of 100 ml of sterile water into each of the culture bottles and the spores were gently dislodged with a clean metal spatula. The spore suspensions were filtered through two layers of cheesecloth and dispensed into a flask. The spore concentration of the two isolates on each of the three substrates was determined with the aid of a hemacytometer. The best substrate for mass production was chosen on the basis of the number of spores per ml of the spore suspensions obtained from each of the grain cultures. Three more 50

trials were carried out with the chosen substrate to check the consistency of spore production. n . . ! ;

... * *

Determination of Spore Germination as a Measure of Spore Viability

In the following experiments, spore viability was measured in terms of the

percentage of spores that produced germ tubes after 2 h on water agar plates at 27°C.

Spores are considered germinated when the germ tubes are twice the length of the

spore width. The percentage germination was calculated using this equation: (number

of spores with germ tubes/total number of spores counted) x 100.

Effect of Short-Term Storage Methods on Spore Viability

Three methods of spore storage were compared in terms of their effect on

spore viability after a 4-week storage period: storage of spores in 1) water at 10°C, 2)

in an oil emulsion (composed of 16% Sunspray 6E and 10% light mineral oil) at

10°C, and 3) on rye grains in vials kept inside a desiccator at room temperature (25 +

3°C). For the first and second methods, spores were harvested from rye grains, pooled together, and suspended in 25-ml of sterile water or oil emulsion in 50-ml flasks. For the third method, 3 grams of fungus-infested rye grains were kept in vials.

The flasks that contained spores in water or oil emulsion were stored at 10°C and the vials that contained the rye grains were stored in a desiccator. Each flask or vial was sampled four times to determine the average percentage germination of the spores per storage method. 51

Germination was determined before and after the 4-week-storage period.

Spores stored in water and oil emulsion were already in suspension; 30 |il droplets of

these suspensions were deposited into each of the four quadrants of a water agar

plate. The fungus-infested grains (5 grains) stored in a desiccator were suspended in

5 ml sterile water and agitated to recover the spores. The resulting suspension was

plated on water agar (30 |il/quadrant). The effect of the storage methods on spore

viability was determined by the difference between the final germination (%

germination at 4 weeks) and the initial germination (i.e. of freshly collected spores,

before storage). ^

Dry Spore Production

Mass-produced fungal spores should be held in a stable and viable state to

ensure long shelf life. Spores of B. sacchari and D. gigantea stored in water or in oil

emulsion for 4 weeks did not lose viability significantly. However, spores stored in

these liquids became contaminated with bacteria and required considerable storage

space. Because of these reasons, it was necessary to devise a method to dry spores

without decreasing spore quality.

Accordingly, the isolates were produced on rye grains, with a modified

methodology. Rye grains were washed and boiled for 10 min, drained, weighed, and

dispensed into glass bottles (50 g/bottle). The bottled grains were autoclaved at

1 2 rc, with 1 5 psi, for 40 minutes for two consecutive days. After cooling, the rye

grains were inoculated with two pieces of 12-mm discs from sporulating 5- to 7-day- old B. sacchari and D. gigantea cultures grown on V-8 agar medium. Inoculated 52

bottles were incubated inside plastic containers at room temperature (25 + 3°C) for

12-14 days. Spores were harvested by the addition of sterile water into the bottles and dislodged from the grain surface with the aid of a sterile metal spatula. The spore suspensions were filtered through two layers of cheesecloth to remove debris and then

through a Whatman filter paper no. 1 (5-cm diameter) to collect the spores for drying.

Whatman filter papers with spores were kept inside plastic petri dishes (15 x 150 nmi) and dried gradually for 5 days at 30°C or 40°C. After five days, the dry spores were scraped from the filter paper and transferred to clean vials for storage at 10°C. Spore germination was determined before and after the spores were dried. The effect of the drying process on spore quality was based on the change in germination, which was

computed by subtracting the initial germination (i.e., of freshly collected, wet spores) value from the final germination value of dried spores.

Effect of Relative Humidity During Storage on Viability of Dry Spores

To create different humidity levels, saturated solutions of NaOH, MnCLa, and

K2SO4 (approximately 100 ml) were prepared and dispensed in small plastic cups according to the methodology described by Winston and Bates (1960). The cups were kept inside 2.02 m^ airtight Rubbermaid®(Rubbermaid Incorporated, Wooster, OH) containers to achieve approximately 30%, 60%, and 90% relative humidity (RH), respectively, at 25°C. Dry spores were dispensed into Eppendorf tubes held upright by Styrofoam tube holders. Three tubes per isolate were placed in each of the humidity chambers along with a data logger (HOBO® weather station, model H08-

004-02; Onset Computer Corporation, Pocasset, MA) to record the temperature and 53

relative humidity during the study. The tubes were kept uncapped to expose the spores to ambient RH. The effect of storage RH on the spore viability was determined from the percent germination of the spore samples every 2 wk for a 10- week period.

The experiment was performed twice with three replications per isolate for each RH. A loopful of spores from each sample was suspended in sterile waster and a 50-|il droplet was spread on water agar plates and the percent germination was determined from three microscopic fields per replicate and averaged. Percent germination was determined after a 2-h incubation on water agar at 27°C with the equation: (number of germinated spores/total number of spores counted) x 100. Fifty spores were counted per replicate.

Effect of Storage Temperature on Viability of Dry Spore

Eppendorf tubes that contained dry spores of the isolates were placed in glass beakers and stored for 12 weeks in three separate incubators at 10°, 25°, and 30°C,

The effect of storage temperature on spore viability was determined from the percent germination values of the samples. Spore germination was checked every two weeks.

A loopful of spores from each sample was suspended in sterile water and a 50-pl droplet was spread on water agar plates and the percent germination was determined from all the spores in three microscopic fields per replicate (minimum of 50 spores/treatment). Percent germination was determined after a 2-h incubation on water agar at 27°C with the equation: (number of germinated spores/ total number of 54

spores counted) x 100. Fifty spores were counted per replicate. The experiment was performed twice with three replications per isolate per temperature level.

Statistical Analysis

Percentage values were transformed using an arcsine square root transformation. The analysis of variance (ANOVA) tests were done using the transformed values and the means were compared by the Fisher's Protected LSD test.

A significance level of P=0.05 was used for the mean comparisons. A regression

analysis was performed to determine the effect of temperature and relative humidity on the germination of stored conidia over time.

Results

Evaluation of Grains as Substrates for Mass-Production of Spores

In the first trial, the spore production by B. sacchari on the three substrates ranged from 1.5-5.2 x 10^ spores per ml. The number of spores of B. sacchari produced on oat, rye, and sorghum grains did not differ from one another (Fig. 3-1

A). On the contrary, the number of spores produced by D. gigantea was significantly higher on rye (2.7 x 10^ spores/ml) than on oats (4.3 x 10^ spores/ml) and sorghum

(2.2 X 10^ spores/ml) (P=0.05, LSD).

In the second trial, B. sacchari produced more spores on rye (1.09 x 10^ spores/ml) than on oats (1.8 x 10^ spores/ml) and sorghum (2.7 x 10^ spores/ml)

(P=0.05, LSD). Again, D. gigantea produced the most spores on rye (1.67 x 10^ spores/ml) than the other two substrates (P=0.05, LSD). There was a significant 55

isolate and substrate interaction for both trials (Table 3-1). Drechslera gigantea produced more spores than B. sacchari on rye (Table 3-2). On the basis of these results, rye was chosen as the substrate for the mass production of spores of B. sacchari and D. gigantea for greenhouse and field experiments.

The isolates produced black, powdery spores on the rye grain after 14 d (Fig.

3-2). The average spore yields for fi. sacchari was 1.46 x 10^ spores/ml and 1.73 x

10^ spores/ml for D. gigantea.

Effect of Storage Methods on Spore Viability

The isolate and the storage method had significant effects on spore

germination, but the trial and the interaction between isolate and storage method had

no effect on spore germination (Table 3-2). Among the three storage methods,

storage of spores on the rye grains in a desiccator had the most significant reduction

in spore germination (P=0.05). Germination of spores of B. sacchari and D. gigantea

was reduced from 99.63% to <2% after 4 wk of storage on the rye grains in a

desiccator. Spores of B. sacchari stored in water or oil at 10°C had >98% germination

before and after storage. Percent germination of spores D. gigantea stored at 10°C in

water was reduced from 99.38% to 94.61% and from 99.48% to 97.51%, in oil.

There was less reduction in the germination of spores of B. sacchari as compared to

spores of D. gigantea. 56

A

B. sacchari D. gigantea

Isolate

B

B. sacchari D. gigantea

Isolate

Fig. 3-1. Production of spores of Bipolaris sacchari and Drechslera gigantea on

solid substrates (grains) after a 14-d incubation at 27° C. Substrates (25 g) in trial 1 (A) were inoculated with 5 ml of blended mycelium and with 10 ml blended mycelium in trial 2 (B). 57

Table 3-1. Analysis of variance for spore production of B. sacchari and D. gigantea on solid substrates (grains) SS MS F Pr>F value

1 nai 1

1 1 ^^080999^19 Isolate I i .OUoUZZZrl 1 L 1 .OWoWZZZJj> 1 z tZ. I\J

Substrate L Z. 1 UJtOO 7Xj 1 z isolate A L 26 64 <0 0001 substrate

Trial 2

Isolate 1 505012500000 505012500000 16.50 0.0016 Substrate 2 4.4917194E12 2.2458597E12 73.39 <0.0001 Isolate X 2 387408333333 193704166667 6.33 0.0133 substrate

DF = degrees of freedom. SS = sum of squares. MS = mean square error. 58

Fig. 3-2. The system used for mass production of Bipolaris sacchari and Drechslera gigantea spores using a solid-substrate (grain). Top photo: autoclaved rye grain in bottles that have been inoculated with the two isolates. Bottom photo: rye grain covered with black, powdery spores of B. sacchari after a 14-d incubation in the dark at 27°C. 59

Effect of Drying on Spore Viability

The drying did not have an adverse effect on the spores of B. sacchari and D. gigantea. For both isolates, the percent germination of the spores was >98% before and after drying at 30° or 40°C.

Effect of Relative Humidity During Storage on Viability of Dry Spores

In general, a significant reduction in spore germination was observed when spores were stored for 10 weeks at high humidity (90%) (P=0.05). In the first trial, only humidity had a significant effect on the germination of stored spores of B. sacchari and D. gigantea (Table 3-3). Based on the slopes of percent germination, spores stored at 90% humidity had reduced germinability over time, whereas those stored at 30% and 60% humidity were not affected. In the second trial, both isolate type and humidity had significant effects on spore germination (Table 3-3).

Germination of spores of B. sacchari was significantly lower at all levels of humidity as compared to spores of D. gigantea.

Similar to the results of the first trial, the germination of spores of B. sacchari and D. gigantea stored at 30% and 60% humidity were not reduced over time, whereas the spores stored at 90% humidity germinated at significantly reduced percentages over time (Fig. 3-3). Spores stored for 10 weeks at 90% humidity had

<32% germination. Spores stored at 30% and 60% had >90% germination after the

10-week storage period. 60

Table 3-2. Analysis of variance for spore germination of B. sacchari and D. gigantea after storage in water, oil emulsion, and on rye grain Source DF SS MS F value Pr>F

Isolate 2 21.41179497 10.70589749 883.96 <0.0001

Storage method 1 0.13088152 0.13088152 10.81 0.0021

Trial 1 0.00455395 0.00455395 0.38 0.5431 Isolate X storage 2 0.02862877 0.01431439 1.18 0.3169 method

DF = degrees of freedom. SS = sum of squares. MS = mean square error. 61

Table 3-3. Analysis of variance for spore germination of B. sacchari and D. gigantea Source DF SS MS F value Pr > F

Trial 1 Humidity 2 0.10218847 0.05109424 398.54 <0.0001

Isolate 1 0.00023281 0.00023281 1.82 0.2027 Humidity X isolate 2 0.00007650 0.00003825 0.30 0.7474

Trial 2 Humidity 2 0.06861419 0.03430710 1842.88 <0.0001

Isolate 1 0.00137008 0.00137008 73.60 <0.0001 Humidity x isolate 2 0.00013341 0.00006671 3.58 0.0602

DF= degrees of freedom SS = sum of squares MS = mean square error Humidity = relative humidity (RH) at which the spores were stored. 62

Table 3-4. Analysis of variance for spore germination of B. sacchari and D. gigantea Source DF SS MS F value Pr > F

Trial 1 Temperature 2 0.00006462 0.00003231 0.51 0.6113

Isolate 1 0.00012663 0.00012663 2.01 0.1817 Temperature X isolate 2 0.00000003 0.00000001 0.00 0.9998

Trial 2 Temperature ''2^ 0.00161900 0.00080950 10.99 0.0019

Isolate 1 0.00001131 0.00001131 0.15 0.7021 Temperature X isolate 2 0.00000507 0.00000253 0.03 0.9663

DF= degrees of freedom. SS = sum of squares. MS = mean square error. 63

Fig. 3-3. The relationship of relative humidity (% RH) during storage and percent spore germination of B. sacchari (o and broken line) and D. gigantea (• and solid line) after storage for 10 weeks at 25°C. Results from trial 1 (A) and trial 2 (B) are presented. 64

o 0.007 o = -5E-05x2+0.002x--5E-05x2- 0.016 " y J.-*' FPFf = 1 a o -0.003

-0.013 y = -5E-05X2 + 0.0021X - 0.0217 0) 5 = c R2 = 1 S i -0.023 - (0 o>

0) -0.033 o 0 10 20 30 40 (0 Storage temperature (C)

Fig. 3-4. The effect of storage temperature on the percentage of spore germination of B. sacchari (o and broken Hne) and D. gigantea (• and solid line) after storage for 12 weeks. Results of trial 1 (A) and trial 2 (B) are presented. 65

Effect of Storage Temperature on Viability of Dry Spores

In the first trial, storage temperature did not have a significant effect on spore germination (Table 3-4; Fig. 3-4 A). The average germination of spores of B. sacchari and D. gigantea stored at the three temperatures was 98.36%. However, in

the second trial, temperature had a significant effect on spore germination (Table 3-

4). Spores stored at 30°C germinated at reduced levels over time (Fig. 3-4 B), whereas those spores stored at lower temperatures were not similarly affected. After

12 weeks of storage, the percentage of germination of spores of B. sacchari stored at

10°, 25°, and 30°C was 97.6%, 96.6%, and 92.04%, respectively. Percentage of germination of spores of D. gigantea stored at 10°, 25°, and 30°C for 12 weeks was

95.09%, 94.7%, and 89.98%, respectively.

Discussion

Production of spores through simple, inexpensive, and reliable methods is an

important aspect of the development of bioherbicides because studies in the

greenhouse and field require a steady supply of ample and viable inoculum. In this

study, mass production of conidia of B. sacchari and D. gigantea was accomplished

in the laboratory through solid-substrate culturing with three species of grain (oat,

rye, and sorghum) as culture media. Of the grains tested, higher spore production

(10^ spores/ml) occurred on rye grain.

One of the problems associated with solid-substrate fermentation is the

recovery of the spores from the substrate (Boyette et al., 1991; Churchill, 1982). In

some cases, the spores are not recovered from the culture substrate; rather, the 66

substrate is incorporated in the final formulated product (Greaves et al., 1998). Abbas and Boyette (2000) applied spores of Colletotrichum truncatum (Schwein.) Andrus &

Moore, a pre-emergence mycoherbicide for hemp sesbania (Sesbania exaltata [Raf.]

Rydb. ex A. W. Hill), with the rice, com, and sorghum grains that were used to produce them. Hildebrand and McCain (1978) used wheat straw infested with

Fusarium oxysporum Schlechtend.: Fr f.sp. cannabis Noviello & W. C. Snyder to control illegal crops of marijuana {Cannabis sativa L.). Oat seeds infested with F.

solani (Mart.) Sacc. f. sp. cucurbitae W. C. Snyder & H. N. Hans were used by

Boyette (1982) to control Texas gourd (Cucurbita texana Gray). The disadvantages of not separating the spores from the culture media include the difficulty in quantification of the fungus, maintenance of sterility of stored inoculum, and the

storage of the bulky solid formulation until its use (Boyette et al., 1991).

Formulation of spores in solid substrate may not be an option for fungi that

will be applied using conventional sprayers. In this study, the storage of spores in the

solid substrate reduced spore viability even after a short-term storage. For both

isolates, short-term storage of spores with the substrate (rye grain) in a desiccator at

room temperature significantly reduced spore quality, as evidenced by the very low

spore germination. One of the possible reasons for the reduction in spore quality after

harvest is that the moisture content of the spores before storage was not optimal to

prolong viability. Another possible reason is that the conditions inside the desiccator

were not favorable to maintain spore viability or to prevent the deterioration of the

substrate and loss of spore quality. 67

Spores of fungal biocontrol agents should be amenable to long-term storage as dry preparations to ensure good shelf life (Jackson, 1997). Fungal spores may begin to deteriorate right after harvest if the moisture content is not favorable (Burges and

Jones, 1998). If the spores are dried before storage, it may prevent them from

deterioration because drying reduces the metabolic activity of the spores. Reduced

metabolic activity minimizes the loss of spore storage reserves and the production of

toxic metabolites (Moore et al., 1996). Li a study by Aregger (1992), Beauveria

hrongniartii (Sacc.) Fetch that was cultured on barley grains and stored with the

grains at 2°C for 24 months had a decline in spore viability over time. The reduction

in spore viability and efficacy of B. brongniartii on white grubs (Melolontha

melolontha L.) was attributed to the continued growth of the fungus, which may have

depleted the spore storage reserves, on barley grain during storage at 2°C.

There are a number of methods to dry spores of various fungi. These methods

include freeze-drying, air-drying, vacuum-tumble drying, and various combinations

of these methods. A method that is sufficiently mild for one type of fungus may

totally destroy the viability of another (Churchill, 1982). Tolerance to desiccation

allows the spores to be dried and stored in a stable form without any significant

reduction in viability. Tolerance to desiccation may also allow spores to survive dry

conditions in the field once the spores have been applied to the target weed. In this

study, spores of B. sacchari and D. gigantea were not adversely affected by drying at

" • 30°or40°C. { f) . J

Storage of dry spores at low humidity and low temperature maintained the

viability of dry spores. Storage of spores at 25°C with 30% and 60 % humidity for 10 68

weeks did not result in a significant reduction in spore germination. Similarly,

storage at 10° and 25°C with approximately 30% humidity did not reduce spore

germination after 12 weeks. Storage of spores at high relative humidity and high

temperature was detrimental to the viability of dry spores of B. sacchari and D.

gigantea.

Generally, fungal spores survive longer in low humidity and low temperature

storage, as shown by several studies. In a study by Abbas and Boyette (2000),

Colletotrichum truncatum-micsXeA rice stored for 6,12, and 24 months at 22-24°C

caused respectively 67-93 %, 39-81%, and < 2% mortality in plants of hemp sesbania.

However, C. truncatum-\vd&si&di rice stored at 4-6°C retained good efficacy for 24

months, and when frozen, for 8 years (Abbas and Boyette, 2000). Spores of the

entomopathogenic fungi Beauveria bassiana (Bals.) Vuill. and Metarhizium

anisopilae (Metsch.) Sorok. stored under these conditions survived for 2 years (Clerk

and Madelin, 1965; Daoust and Roberts, 1983; Steinhaus, 1960). Dried spores of M. flavoviridae Gams and Rozsypal, a biocontrol agent against desert locusts and

grasshoppers, had 95% germination when stored as powder 10°-14°C compared with

only 27% when stored at 28°-32°C (Moore et al., 1996). Two isolates of C. truncatum

showed only minimal decay in spore viability after storage at low temperatures (4°

and 15°C) for 58 days (Silman et al., 1993).

Grain as substrates was suitable for mass-production of spores of B. sacchari

and D. gigantea. There are many other inexpensive and readily available natural

substrates that can be utilized in the mass-production of inoculum of other promising

biocontrol agents. The results from this study allowed for the mass-production and 69

storage of inoculum of B. sacchari and D. gigantea without loss of viability until

inoculations in the greenhouse or field were performed. It is recommended that experiments with narrower temperature and relative humidity intervals be performed to determine the optimum storage temperature and relative humidity for each isolate. CHAPTER 4

THE EFFECT OF OIL FORMULATION ON THE BIOHERBICIDAL EFHCACY OF Bipolaris sacchari AND Drechslera gigantea

Introduction

Formulation in the case of a bioherbicide refers to the blending of the active ingredient, in the form of living propagules, with a carrier or solvent and typically other adjuvants to produce a final product that can be delivered effectively to the

target weed (Boyette et al., 1991; Rhodes, 1993). The requirement for appropriate formulations that will store well and promote successful establishment of the pathogen in the field are two of the major challenges to the development and marketing of bioherbicides (Auld, 1992; Baker and Henis, 1990; Greaves and

Macqueen, 1990; Rhodes, 1993).

Formulation attempts to overcome the constraints imposed by the target weed and the environment on the bioherbicide through enhanced biocontrol efficacy and ease of application. Formulation can stimulate spore germination (Winder and Van

Dyke, 1990), improve virulence of the agent (Bothast et al., 1993), allow adequate spread of the inoculum on leaf surfaces, prevent wash-off by rain, protect against ultraviolet radiation, and improve the ease of application with traditional spray equipment (Greaves and Macqueen, 1990). Formulation can overcome the requirement for extended dew periods or for free moisture for infection (Auld, 1992;

Connick et al., 1990). Also, formulation can reduce the inoculum threshold of the

70 71

bioherbicidal agent needed to enhance its stability, increase its biological activity,

interfere with the host defenses, and affect the plant cuticle (Amsellem et al., 1991;

Auld, 1992; Boyette et al., 1991; Burge, 1988). Formulations should be safe, easy to use, and maintain the viability of the biological propagules during storage,

distribution, and application (Leggett and Gleddie, 1995; Rhodes et al., 1990).

Additives that may be used in bioherbicide formulation include surfactants,

humectants, and stickers. Surfactants are used to ensure uniform distribution of the

inoculum on the leaf surfaces. They can penetrate the cuticle and aid in the retention

of the inoculum, thereby the infection process is enhanced. Surfactants have been

widely used in formulations although several have been found to be toxic to some

fungi. Humectants are used to stabilize the moisture content of the bioherbicide

regardless of fluctuations in humidity. However, the cost and toxicity of humectants

have deterred their wide use in bioherbicidal formulations. Stickers increase the

adhesion of spores to the leaf surface and prevent the spores from being washed off

by rain. Some stickers can also serve as humectants.

Invert emulsions have been used in bioherbicide formulations. An invert

emulsion is composed of water surrounded by a layer of oil. It can hold propagules in

the water phase protected within the oily matrix. The oil improves the adhesion of

the propagule on the host surface, while the water phase allows the spores to

germinate (Boyette et al., 1993; Connick et al., 1991; Daigle et al., 1990; Munyaradzi

et al., 1990; Womack and Burge, 1993; Womack et al., 1996). Altemaria cassiae

Jurair & Khan formulated in an invert emulsion composed of lecithin, paraffin oil,

and wax gave >95% control of sicklepod (Senna obtusifolia [L.] Irwin & Bameby), 72

with little or no dew, in trials in the greenhouse and field (Quimby et al., 1988). The invert emulsion also enhanced the field performance of Colletotrichum truncatum

(Schwein.) Andrus & Moore against hemp sesbania {Sesbania exaltata [Raf.] Rydb. ex Hill). The fungus-invert emulsion formulation controlled 95% of the target weed; the fungus without the invert emulsion controlled only 10% of the target weed

(Boyette et al., 1993). In preliminary studies with invert emulsion formulation

(Quimby, 1988), the dew requirement was reduced for Ascochyta pteridis Bres., a biocontrol agent for bracken (Pteridium aquilinum [L.] Kuhn) (Womack and Burge,

1993).

Invert emulsions are not widely used because they are complex and are not easy to prepare. They can be highly viscous and hence incompatible with conventional spray equipment. They are generally phytotoxic to plants (Auld, 1993;

Boyette, 1994; Womack, 1993). Because of their ability to promote phytotoxicity, attention has been focused recently on oil emulsions as carriers for microbial herbicides.

The objectives of this study are: 1) to determine whether formulation with oil emulsion can improve the efficacy of B. sacchari and D. gigantea as control agents for cogongrass; 2) to assess the toxicity of the oil emulsion on B. sacchari and D. gigantea spores during storage and application on cogongrass; and 3) to evaluate the suitability of vegetable oils as carriers for B. sacchari and D. gigantea. 73

Materials and Methods

Propagation of Test Plants

Rhizomes were collected from a cogongrass patch near Lake Alice, University of Florida Campus, Gainesville, FL. Cogongrass plants were propagated in the greenhouse from 5-cm rhizome pieces planted in a commercial potting medium

(Metromix 300; Scott' s-Sierra Horticultural Products Co., Marysville, OH) in plastic trays (13 x 17 x 56 cm). Rhizomes with shoots were then transplanted into commercial potting medium in plastic pots (15-cm diameter; 17.5-cm height). The plants were watered regularly and fertilized with Osmocote® 14-14-14 N-P-K controlled-release fertilizer (Scott's-Sierra Horticultural Products Co., Marysville,

OH).

Preparation of Spore Suspensions

Spores of B. sacchari and D. gigantea were obtained from 7- to 10-day-old

V8-juice cultures incubated in the dark at 27°C or from 12- to 14-day-old-rye grain cultures that were incubated at room temperature. The resulting spore suspensions were filtered through two layers of cheesecloth and adjusted to the desired concentration with the aid of a hemacytometer.

Disease Assessment

Disease severity (DS) was determined with the aid of a visual assessment key used for southern com leaf blight (James, 1971), with 50% as the maximum disease

severity. Disease severity per pot was computed by the average of ratings of all the

plants in the pot. Disease severity per plant was assessed per leaf, and all the ratings 74

of all the leaves were added and the sum divided by the total number of leaves of the plant.

Determination of Spore Germination

Spores were observed for the presence of germ tubes after a 2-h incubation at

27°C. Spores are considered germinated when the germ tubes are twice the length of the spore width. The percentage of spore germination was computed by dividing the number of germinated spores by total number of spores counted and multiplying the quotient by 100.

Data Analysis

Data were transformed using an arcsine square root transformation and analyzed using SAS (SAS Institute, Gary, NC). Data from trials of similar

experiments were pooled unless there were significant trial differences (i.e., heterogeneity of variance).

Evaluation of Oil Emulsion as a Bioherbicide Carrier in the Greenhouse

Spores of B. sacchari were formulated with gelatin (Knox® unflavored gelatin; Nabisco, Inc., East Hanover, NJ) and oil emulsions that contained different proportions of Sunspray® 6E (Sun Company Inc., Philadelphia, PA) and light mineral oil (Fisher Scientific, Fair Lawn, NJ). Two oil mixtures were prepared, one composed of 80% Sunspray® 6E and 20% mineral oil, the other containing 8%

Sunspray® 6E and 20% mineral oil. The oil mixtures were added to equal volumes of water with the spores. This resulted in formulations with final oil concentrations of 75

50% and 14% oil. Double-strength gelatin (2%) was prepared and mixed with an equal volume of spore suspension. This resulted in a bioherbicide formulation with

1 % gelatin as the carrier.

With the use of hand-pumped sprayers, 4-week-old plants (five plants per pot; three pots per treatment) were inoculated with the three formulations until run-off.

Plants were also sprayed with the carriers alone, to determine and compare the potential phytotoxicity of gelatin, high oil, and low oil carriers. The experiment was done twice; the B. sacchari spore concentration for the first trial was 4x10^

spores/ml and 3.0 x 10^ spores/ml for the second trial. All treated plants were incubated in a dew chamber (28 + 1°C, 100% RH), arranged in a completely randomized manner. After the 24-h dew period, the plants were transferred to the greenhouse for further observation. Disease severity (DS) and weed mortality (WM) were assessed at 21 days after inoculation (DAI).

Assessment of the Fungitoxicity of Oil Emulsions

Effects of formulation and short-term storage of spores (conidia) with

Sunspray® 6E and mineral oil emulsion on spore viability and spore germination were determined by experiments in vitro (water agar) and in vivo (cogongrass leaves).

Spores of B. sacchari and D. gigantea were mixed with oil emulsions that contained

various amounts of Sunspray® 6E (4, 8,16, and 32%), 10% mineral oil, and water to make up the remainder of the 100%. Spores of B. sacchari and D. gigantea were formulated with the oil emulsions and stored at 10°C for 96 h, with spores suspended in only water to serve as the control. After 24, 48, 72, and 96 h of incubation, the 76

percentage of spore germination was determined by four 30 |il droplets of the spore and oil emulsion mixture deposited on water agar plates (100 x 15 mm petri dish) that had been divided into quadrants. Percent germination on water agar was determined after incubation for 2 h at 27°C. Data from two trials were pooled and analyzed.

Cogongrass plants were inoculated with spores of B. sacchari and D. gigantea formulated with 26% oil (16% Sunspray® 6E + 10 % mineral oil) until complete coverage of the foliage was achieved. Inoculated plants were incubated in the dew chamber (1(X)% RH; 28 + TC) and leaves were taken from the inoculated plants at 2,

4, and 6 h after incubation. These leaves were cut into 2x8 mm pieces and the leaf pieces were placed inside a glass petri dish lined with a filter paper that had been soaked with a 1:1 (v/v) solution of glacial acetic acid and 95% ethanol (Fernando et

al., 1993). The tissues were cleared for at least 24 h to remove the green pigment and make the spores more visible. The samples were stained with acid-fuschin solution, fixed with polyvinyl alcohol (Salmon, 1954), and checked for germinated spores under 200 x and 400 x magnifications.

Efficacy of B. sacchari and D. gigantea Oil Emulsion Formulations under Different Dew Periods

Four-week-old plants of cogongrass (3 plants per pot; 3 pots per treatment) in

clay pots (1 1.5 cm diameter; 1 1.0 cm height) or square plastic pots (8x8x9 cm) were inoculated with spores of B. sacchari and D. gigantea formulated with 14% oil emulsion (4% Sunspray® 6E + 10% mineral oil) as the carrier. Plants were sprayed with hand-pumped sprayers until the foliage was fully covered with the emulsion. 77

For comparison, another set of plants was inoculated with spores formulated with 1% gelatin as the carrier.

Inoculated plants were randomly arranged and incubated in a dew chamber

(100% RH; 28 + TC) for 4, 8, 12, and 24 h or kept in the greenhouse immediately after inoculation (0 h dew period). Disease severity was assessed at 7 and 21 DAI.

The experiments were done twice for each isolate. The spore concentration of the

isolates for the trials was 5x10^ spores/ml.

Evaluation of Various Oils as Bioherbicide Carriers in the Laboratory and Greenhouse

Tests were performed in the laboratory and greenhouse to determine whether some commonly available vegetable oils would be comparable in efficacy to the

Sunspray® 6E + mineral oil formulation. The mineral oil portion of the oil phase was substituted with canola, citrus, com, soybean, and sunflower oils. Ten ml spore suspensions of B. sacchari or D. gigantea that contained 5x10^ spores/ml and 10 ml oil emulsions with Sunspray® 6E (16%) + vegetable oils (20%) were added together to achieve a final spore-oil emulsion formulation with 18% total oil content (8%

Sunspray® 6Eand 10% vegetable oil). Spores in water only and spores in Sunspray®

6E + mineral oil + water were also prepared for comparison. Percent germination was

determined by depositing four 30 |il droplets of the spore and oil emulsion mixture on water-agar plates (100 x 15 mm petri dish) that had been partitioned into quadrants.

The spores were incubated on water agar plates for 2 h at 27°C. After incubation, the percentage germination was determined with the previously described methodology.

Percent germination of spore was determined before and after storage in the various 78

oil emulsions for 24, 48, 72, and 96 h at 10°C. In the first trial, com, citrus, soybean,

and sunflower oils were evaluated. In the second trial, canola oil was evaluated along

with the other oils, while citrus oil was excluded because it was fungitoxic in the first

trial.

The performance of the various oil emulsions as bioherbicidal carriers was compared in greenhouse tests with spores of B. sacchari as the active ingredient.

Spores in vegetable-oil emulsions were prepared by mixing together equal volumes of spores of B. sacchari in water and an oil emulsion composed of 18% vegetable oil

(canola, com, sunflower, or soybean), 2% emulsifier (Myverol®18-99; Eastman

Chemical Company, Kingsport, TN), and 80% sterile water. Spores in petroleum- based oil emulsion were prepared by mixing equal volumes of spores in sterile water and a mixture of 8% Sunspray® 6E, 10% mineral oil, and 82% sterilized water. The resulting spore-oil emulsion formulation was composed of either 9% vegetable oil or

Sunspray® 6E + mineral oil and had a spore concentration of 5 x 10^ spores/ml.

Cogongrass plants in plastic pots (10 cm diameter; 8.8 cm height) were inoculated with the spores formulated with the various oils. Plants were sprayed until the surface of the foliage was fully covered with the emulsion with hand-pumped sprayers and incubated in the dew chamber for 12 h (100% RH; 28 + TC). After incubation, the plants were moved to a greenhouse for further observations and assessment of disease severity. 79

Results

Evaluation of Oil Emulsion as a Bioherbicide Carrier

Spores of Bipolaris sacchari formulated with oil emulsions that contained

14% (low oil) and 50% (high oil) Sunspray® 6E + mineral oil caused significantly higher disease severity (DS) levels and weed mortality (WM) than spores formulated with 1% gelatin (Fig. 4-1, A and B). Plants inoculated with the spores in gelatin formulation exhibited mostly discrete and some coalesced lesions along the leaf blade and blight of leaf tips. Plants sprayed with spores in the oil emulsions exhibited severe blight of the foliage and plant death. Gelatin alone had no phytotoxic effects on cogongrass (0% WM), whereas the oil emulsion with 50% oil was significantly more phytotoxic than the emulsion with 14% oil (Fig. 4-1 C). However, when mixed with spores of B. sacchari, both emulsions caused similar severity of disease and weed mortality. Thus, with the same inoculum level, similar levels of disease severity and weed mortality could be achieved with the lesser amount of oil.

Evaluation of Fungitoxicity of the Oil-Emulsion Carrier

Spores stored for 24, 28, 72, and 96 h in water (0% oil) or oil emulsions that contained 18%, 26%, and 42% Sunspray® 6E + mineral oil germinated on water agar

at 27°C 2 h after plating. Isolate type and storage time in water or oil emulsion

significantly affected spore germination (Table 4-1). The average germination of

spores stored for up to 96 h in water and oil emulsions was 93.4% for B. sacchari and

97.5% for D. gigantea (Fig. 4-2). Overall, spores of D. gigantea had a higher

percentage germination than spores of B. sacchari. Spore germination was slightly 80

lower after storage for 72 h (94.5%) and 96 h (92.2%), but still >90% (Fig. 4-2).

Spores formulated with the oil emulsion produced germ tubes on cogongrass leaf tissues within 2-6 h after inoculation and incubation in a dew chamber (Fig. 4-3).

Efficacy of B. sacchari in Oil-Emulsion Formulation under Different Dew Periods

In the first trial, the formulation, length of dew exposure (dew period), and the interaction of formulation and dew period significantly influenced the disease severity caused by B. sacchari on cogongrass (Table 4-2). Plants inoculated with spores formulated with the oil emulsion exhibited severe blight and plant death irrespective

of the dew period (0, 4, 8, 12, or 24 h). In the second trial, formulation and dew period had significant effects on DS but there was no significant interaction effect

(Table 4-2). Spores formulated with oil emulsion caused higher DS than spores

formulated with gelatin over all dew periods tested (Figs. 4-4 and 4-5). With both

formulations, the DS level increased with prolonged dew exposure immediately after

inoculation (Fig. 4-5).

Efficacy of D. gigantea in Oil-Emulsion Formulation under Different Dew Periods

Although similar results were obtained from both trials with D. gigantea, the

level of variance between the trials did not allow the data to be combined for analysis.

Formulation and length of dew exposure significantly influenced DS caused by D.

gigantea (Table 4-3). Spores formulated in the oil-emulsion caused significantly

higher DS with any length of dew exposure after inoculation as compared to spores 81

60 A A 5* 50 40 > B « 30 o 20 CO CO 10 o « 0

Spores + gelatin Spores + low oil Spores + high

oil

Formulation

B

gS 120 A A 100 n 80 r o 60 E 40 20 B 0

Spores + Spores + low oil Spores + high

gelatin . oil

Formulation

120 100 2* 80 tCO c 60 E 40 o H V 20 0) 0

Gelatin Low oil High oil

Carrier

Fig. 4-1. Effect of gelatin and oil formulations on the biocontrol efficacy of Bipolaris sacchari on cogongrass at 21 DAI (A and B). The effect of carriers alone (control) on cogongrass (C). In each graph, bars with different letters are significantly different at P=0.05 as determined by Duncan's Multiple Range test. 82

Table 4-1. Analysis of variance for spore germination of B. sacchari and D. gigantea Source DF SS MS F value Pr>F

Isolate 1 0.74553882 0.74553882 36.36 <0.0001 Oil concentration 3 0.08876085 0.02958695 1.44 0.2310 Storage time 3 0.56680707 0.18893569 9.22 <0.0001 Isolate X oil concentration 3 0.07843309 0.02614436 1.28 0.2836 Isolate X storage time 3 0.00327712 0.00109237 0.05 0.9838 Oil concentration x 9 0.12486246 0.01387361 0.68 0.7297 storage time

DF = degrees of freedom. SS = sum of squares. MS = mean square error. —

83

80

ra c 60 0% — 18% 0) 40 O) A 26% £ 20 o K 42% Q. 0 24 48 72 96

Storage time in oil emulsion (h)

B

3- 100 c o 80 C 60 -

40 o> £ 20 o a. u) 0 24 48 72 96

Storage time in oil emulsion (h)

Fig. 4-2. Effects of formulations and short-term storage in oil emulsions with increasing oil concentrations on the germination of spores of Bipolaris sacchari (A) and Drechslera gigantea (B). 84

A

Fig. 4-3. Spore germination of Bipolaris sacchari (A) and Drechslera gigantea (B) on cogongrass leaf (magnified 200 x). Photos were taken at 6 h after the plants were inoculated with spores in an emulsion (26% oil) and incubated in a dew chamber at 27°C and 100% RH. 85

Table 4-2. Analysis of variance for disease severity of cogongrass inoculated with B. sacchari Source DF SS MS F value Pr>F

Trial 1

Formulation 1 2.22315842 2.22315842 867.46 <0.0001 Dew period 4 0.52412652 0.13103163 51.13 <0.0001 Formulation X dew period 4 0.51809512 0.12952378 50.54 <0.0001

Trial 2

Formulation 1 0.51993063 0.51993063 34.42 <0.0001 Dew period 4 0.62324390 0.15581097 10.31 <0.0001 Formulation X dew period 4 0.03043727 0.00760932 0.50 0.7331

DF = degrees of freedom. SS = sum of squares. MS = mean square error. 86

Table 4-3. Analysis of variance for disease severity of cogongrass inoculated with D. gigantea Source DF SS MS F value Pr>F

Trial 1 Formulation 1 0.39808383 0.39808383 18.81 <0.0001 Dew period 4 0.53649472 0.13412368 6.62 0.0004 Formulation X dew period 4 0.15540838 0.03885209 1.84 0.1374

Trial 2

Formulation 1 0.17820685 0.17820685 12.05 0.0011 Dew period 4 0.81603353 0.20400838 13.79 <0.0001 Formulation X dew period 4 0.10440448 0.02610112 1.76 0.1508

DF = degrees of freedom. SS = sum of squares. MS = mean square error. 87

Fig. 4-4. Biocontrol efficacy of spores of B. sacchari and D. gigantea formulated with gelatin or oil emulsion and exposed to various lengths of dew after inoculation. Top photo: (L-R) plants inoculated with spores of B. sacchari formulated with 1% gelatin and exposed to 0, 4, 8, 12, and 24 h of dew or formulated with an oil emulsion (4% Sunspray 6E + 10% mineral oil) and exposed to 0, 4, 8, 12, and 24 h of dew. Bottom photo: (L-R) plants inoculated with spores of D. gigantea formulated with

1% gelatin and exposed to 0, 4, 8, 12, and 24 of h dew or formulated with an oil emulsion (4% Sunspray® 6E + 10% mineral oil) and exposed to 0, 4, 8, 12, and 24 h of dew. 88

A

o < 0 0 4 8 12 24

Dew exposure (h)

B

< 0 -I 1 1 1 1 0 4 8 12 24

Dew exposure (h)

Fig. 4-5. Enhancement of biocontrol efficacy of B. sacchari on cogongrass with spores formulated in an oil emulsion (• and broken line) that contained 14% oil (4% Sunspray 6E + 10% mineral oil) compared with formulation in 1% gelatin (o and solid line). Disease severities (transformed) at different dew periods are shown by A

(trial 1) and B (trial 2). 89

A

n Q

0.8 -

0.7 - y = 0.0301x + 0.3798 0.6 - = 0.4352 ^ 0.5 - •

0.4 -

0.3 - ^'^"""''^ 0.2 - y = 0.0921 X - 0.0001 = 0.9446 0.1 -

- 1 0 1 ; 0 4 8 12 24

Dew exposure (h)

B

»o 0.9 ? 0.8 H y = 0.0466X + 0.4989 Q 0.7

0.5 I 0.0999X + 0.2332 £ 0.4 « \ R^ = 0.9472 § 0.3 0) JT 0.2 H I 0-1 H

4 8 12 24

Dew exposure (h)

Fig. 4-6. Enhancement of biocontrol efficacy of D. gigantea on cogongrass with spores formulated in an oil emulsion (• and broken line) that contained 14% oil (4% Sunspray 6E + 10% mineral oil) compared with formulation in 1% gelatin (o and solid line). Disease severities (transformed) at different dew periods are shown by A (trial 1) and B (trial 2). 90

Table 4- 4. Analysis of variance for spore germination of B. sacchari and D. gigantea Source DF SS MS F value Pr>F

Oil type 6 76.76482775 12.79413796 822.74 <0.0001

Isolate 1 0.60179727 0.74008757 47.59 <0.0001 Storage time 4 0.22898369 0.05312873 3.42 0.0092 Oil type X storage time 24 0.24927612 0.01038650 0.67 0.8829 Oil type X isolate 6 0.14352456 0.02392076 1.54 0.1642 Oil type X isolate x storage 28 0.030766350 0.01098798 0.71 0.8671 time

DF = degrees of freedom. SS = sum of squares. MS = mean square error. 91

A

£^ 100 c a a a o 80 a (0 c 60 40 20 o a 0 (A c CO c CO 0) 0) o o (U c c 0) o CO (B O O O o c CO

Formulation

B

50

2 40 - a a a a a a I" 30 >0) 0) (0 20 0> 8 10 H 0) « 0 c o + 2 0) o ^ 2 £ o I « o CO O I P3; 2 o S W + CO + CO + Formulation

Fig. 4-7. (A) The effect of various oil fomulations on the spore germination of B. sacchari and D. gigantea. The bars in A represent the combined germination values of both isolates. Bars with the same letters are not significantly different at P=0.05 as determined by Duncan's Multiple Range test. (B) The efficacy of B. sacchari formulated with various vegetable oils and Myverol or Sunspray® 6E + mineral oil.

Dotted bars represent results from trial 1; white bars represent results from trial 2.

Within the same trial, bars with the same letters are not significantly different at

/'=0.05, as determined by the Duncan's Multiple Range test. 92

formulated with 1% gelatin. Regardless of formulation, the levels of DS were higher when the inoculated plants were exposed to longer dew periods (Figs. 4-4 and 4- 6).

Evaluation of Various Oils as Bioherbicide Carriers in the Laboratory and Greenhouse

Data from two trials were combined and analyzed. Among the various oils

tested, only citrus oil inhibited the germination of spores of B. sacchari and D.

gigantea (Fig. 4-7 A). Spores stored in emulsions that contained 4% Sunspray® 6E

and 10% of one of the other oils (canola, com, sunflower, soybean, and mineral) for

24, 48, 72, and 96 h germinated on water agar as early as after a 2-h incubation at

27°C.

Oil type, isolate type, and duration of storage (storage time) in the oil

emulsions had significant effects on spore germination (P=0.05) (Table 4- 4).

Germination of spores stored in water, com oil, mineral oil, sunflower oil, and

soybean oil ranged from 96.5-97.3% (Fig. 4-7 A). Compared with those stored in

com, mineral, soybean, sunflower, and water, spores formulated with citras oil did

not germinate. In general, spores of B. sacchari had higher percentage germination

(90% on the average) than spores of D. gigantea (87% on the average) (data not

shown). The average germination values for spores stored in oils or water for 0, 24,

48, 72, and 96 h were 89.4%, 87.2%, 89.5%, 87.6%, and 88.1%, respectively.

Germination did not necessarily increase or decrease with time, therefore, the

differences in the germination values are attributed to spore variability rather than to

the effect of prolonging the storage of the spores in the oil emulsions. 93

In terms of the level of disease, the vegetable oil (canola, com, soybean, or sunflower) + Myverol formulations were inferior to the Sunspray® 6E + mineral oil

formulation (Fig. 4-7B). In the first trial, all formulations caused >40% DS. In the second trial, spores formulated in canola oil caused significantly higher DS (47.5%) than the other formulations (< 39%). In both trials, the DS level increased with time;

DS level was higher at 21 DAI than at 7 DAI (data not shown). However, spores formulated with any of the vegetable oils + Myverol tended to clump together, necessitating constant shaking of the inoculum during application.

Discussion

Various additives have been used to improve or modify spore germination, pathogen stability and virulence, environmental requirements, or host preference of

bioherbicidal fungi (Boyette et al., 1996). Among these various additives are invert

(water-in-oil) emulsions and oil (oil-in-water) emulsions. Invert emulsions retard evaporation by trapping water in the spray mixture. They provide the free moisture

required for spore germination and infection (Quimby, 1988; Daigle et al., 1990;

Amsellem et al., 1990). Invert emulsions can also reduce or eliminate the

requirement for high levels of inoculum (Amesellem et al., 1990). Compared to invert emulsions, oil emulsions are easier to prepare, less viscous, and are more compatible with conventional spray equipment. Oil emulsions are suitable bioherbicide carriers because of their properties of surface wetting on the leaf, spore protection, and some water retention (Auld, 1993). Oil emulsions have been used in

several bioherbicide formulations with some success (Greaves et al., 1998). Under controlled environment experiments, Colletotrichum orbiculare (Berk. & Mont.) Arx 94

formulated with a vegetable oil emulsion caused better development of anthracnose on bathurst burr {Xanthium spinosum L.) in the absence of dew as compared to aqueous suspensions (Auld, 1993). Formulation of C. truncatum with unrefined com

oil reduced the minimum dew period requirements for infection of hemp sesbania

(Sesbania exaltatd) and weed mortality from 12 h to 2 h. The enhancement of efficacy of C. truncatum with the oil formulation is likely from the ability of the unrefined com oil emulsion to reduce evaporation and retain water (Boyette, 1994).

Unrefined com oil stimulated conidial germination and appressorial formation of C. truncatum, this resulted in increased infection of hemp sesbania (Egley and Boyette,

1994). Formulation of Altemaria helianthi (Hansf.) Tub. Et Nish., a biocontrol agent for cocklebur {Xanthium strumarium L.), in an emulsion of unrefined com oil allowed the pathogen to infect the target weed, with or without dew at 24 h after inoculation

(Abbas and Egley, 1996). The dew period requirement of Mycocentrospora acerina

(R. Hartig) Deighton for infection of field pansy (Viola arvensis Murray) was reduced from >36 h to 12 h by the inoculum formulated with rapeseed oil-water emulsion (1:9 v/v) (Potyka, 1996). Chandramohan (1999) effectively controlled seven weed species by three pathogens formulated with an oil emulsion composed of Sunspray 6E (40%) and paraffin oil (10%). The oil-emulsion formulation caused significantly higher disease levels as compared to the 0.5% aqueous Metamucil formulation.

An oil emulsion similar to the one used by Chandramohan (1999) was evaluated in this study. In initial experiments in the greenhouse, the Sunspray 6E + mineral oil emulsion, that contained 50% and 14% total oil, improved the level of control of cogongrass. The high level of disease and damage on cogongrass was 95

attributed to the combined action of the active ingredient (spores of B. sacchari) and

the oil emulsion carrier, which was, to some extent, phytotoxic on cogongrass.

The oil emulsion was tested for fungitoxicity to determine its suitability as a carrier and to exclude the possibility that control in the greenhouse was merely due to phytotoxicity of the emulsion. Short-term storage (96 h) of spores of B. sacchari and

D. gigantea in oil emulsions that contained 14-42% oil did not inhibit spore germination on water agar or on cogongrass. Germ tubes were observed 2 h after inoculation of water agar or cogongrass plants. Therefore, the emulsion did not reduce spore viability during storage and application.

The oil emulsion enhanced the biocontrol efficacy under different dew periods. With any length of dew exposure after inoculation, the emulsion formulation caused significantly higher levels of disease than the gelatin formulation. Within 21 days, plants treated with the gelatin formulation exhibited mostly distinct lesions along the leaf blade and only minor blighting of the leaf tips, whereas plants sprayed with the emulsion exhibited severe blighting or plant death. The more uniform the coverage of the leaf surfaces by the emulsion-inoculum combined with the

phytotoxicity of the emulsion itself is considered as the basis for the severe blighting.

With the oil formulation, higher levels of disease were achieved with shorter dew

periods in the greenhouse. Therefore, the oil formulation is ideal for use in the field where dew periods may vary.

Com, canola, soybean, and sunflower oils were not toxic to spores of B. sacchari and D. gigantea and did not inhibit spore germination on water agar plates.

In trials in the greenhouse, spores in the vegetable oil + Myverol formulations and the 96

Sunspray® 6E + mineral oil formulation caused similar disease severities on cogongrass. According to these results, emulsions composed of commonly available

oils + a food-grade emulsifier such as Myverol may be used instead of Sunspray® 6E

+ mineral oil as carriers for bioherbicide to achieve high levels of control. However,

in terms of ease of application, the Sunspray® 6E (which has its own emulsifier) +

mineral oil formulation is more desirable because the spores tended to be more evenly

dispersed in the emulsion and frequent mixing was not required. In the vegetable oil

formulations, the spores tend to aggregate on the oily layer at the surface of the

suspension.

In conclusion, the efficacy of the two potential control agents for cogongrass,

B. sacchari and D. gigantea, can be improved by formulating their spores in an oil

emulsion. The oil formulation caused higher levels of disease and damage even with

limited period of dew after inoculation. With some modifications or with better

emulsifying agents than Myverol, these vegetable oils may be used in formulations

instead of Sunspray® 6E and mineral oil. CHAPTER 5 EVALUATION OF HOST RANGE OF Bipolaris sacchari

Introduction

Host-plant specificity has been regarded as one of the most important factors in the selection of pathogens as biological control agents (Charudattan, 1989; Watson, 1985).

Determination of host range is especially important in the case of nonindigenous agents used

as classical biocontrol agents, because of concerns about damage to nontarget hosts as well

as endangered plants. In the case of indigenous pathogens used as bioherbicides,

determination of host range is important to determine the potential nontarget hosts that may be attacked by the bioherbicide pathogen and to take the necessary precaution, such as

restricted labeling, to avoid injury to nontarget plants. Nontarget plants in this case may be

economically important crops and plants for which there may be potential conflict-of-

interest issues (Weidemann, 1991).

Plant pathogens may have narrow or broad host ranges, but most, whether facultative

saprophytes or obligate parasites, are generally pathogenic to one or few species (Gaiimann,

1950; Holcomb, 1982; Johnson, 1976; Oku et al., 1979). Brian (1976) compared the levels

of specificity of fungal plant pathogens up to the level of formae speciales and concluded

that most fungal pathogens show a high degree of host specificity. While specificity of

bioherbicides is a major factor in relation to their environmental safety, it is a major

drawback from the commercial development standpoint. For situations where several weed

species are present, the use of a bioherbicide that only controls one weed species becomes

97 98

uneconomical to develop and commercialize (Greaves et al., 1998). The multiple pathogen

strategy, wherein a bioherbicide is composed of a mixture of fungal pathogens, has been

used to control several species of weedy grasses (Chandramohan, 1999). However, it is not clear whether a bioherbicide composed of several pathogens will simply multiply the cost of development and therefore prove to be uneconomical. The formulation of fungal pathogens with various additives to enhance their biocontrol efficacy has also resulted in a wider host range (Abbas and Boyette, 1994).

The centrifugal-phylogenetic scheme proposed by Wapshere (1974) has been used extensively by workers in biocontrol of weeds to determine the extent of host range of a biocontrol agent. The scheme includes testing of the other plant species that are

taxonomically (i.e., phylogenetically) related to the weed targeted for control as well as plants that are considered at "risk". Plants considered to be "at risk" include those that are related to the target weed, those that have not been previously exposed to the pathogen, those that have limited information on their natural enemies, those that are similar to the target weed in terms of secondary compounds and morphology, those attacked by related organisms, and those that have been recorded as possible hosts of the candidate agent

(Schroeder, 1983; Wapshere, 1982; Wapshere, 1983). •

Phylogenetic testing is more precise with highly host-specific insects or obligate parasites such as rust fungi. This testing method has been proposed for the determination of host range of biocontrol agents used in the classical approach. For pathogens that are not highly host-specific, such as those commonly used in the bioherbicidal approach, a high

level of weed kill rather than safety to nontarget plants is the prime consideration.

Nonetheless, the host-range tests for bioherbicidal agents should include plants that are 99

ecologically and economically important at the site of application and plants known or suspected to be susceptible to the pathogen (Charudattan, 1989).

The isolate of the fungus Bipolaris sacchari (E.J. Butler) Shoem. is indigenous to

Florida and has been reported to occur on a number of plants, mostly grasses (Alfieri et al.,

1994; Farr et al., 1989). Bipolaris sacchari causes necrotic leaf streak in pampasgrass

{Cortaderia selloana [Schult. & Schult. f.] Asch. & Graebn.), leaf spot in ravennagrass

{Eriathus ravennae [L.] P. Beauv.), and guineagrass (Panicum maximum Jacq.) and eye spot

disease on lemongrass (Cymbopogon citratus [DC. Ex Nees]), napiergrass {Pennisetum purpureum Schumach.), and sugarcane {Saccharum officinarum L.). Bipolaris sacchari is

also a promising biocontrol agent of cogongrass. It causes leaf lesions and leaf blights when

applied inundatively. Cogongrass is an ideal target weed for biocontrol with B. sacchari

because it forms monotypic stands and is primarily a problem in noncrop areas. Therefore,

there is a lower risk of nontarget plant injury.

Drechslera gigantea is another promising biocontrol agent for cogongrass.

Chandramohan (1999) has previously determined the host range of D. gigantea in trials in

the greenhouse. The plant species that were found to be susceptible to D. gigantea are:

bahiagrass, bermudagrass, crowfootgrass, green foxtail, guineagrass, johnsongrass, large

crabgrass, natalgrass, southern sanbbur, Texas panicum, torpedograss, vaseygrass, and

yellow foxtail. In trials in the greenhouse, none of the economically important crop species

tested (36 species, including 53 cultivars) was susceptible to D. gigantea.

The objective of this study is to determine whether economically important crops

and other grass species might be susceptible to Bipolaris sacchari when formulated with oil

emulsion as the carrier. 100

Materials and Methods

Production of Test Plants

The plants tested for their reaction to B. sacchari included several species of cultivated and weedy grasses as well as several economically important plant species that are grown in Florida. A total of 45 plant species in 35 genera and 10 families were evaluated.

Representative species from 10 plant families (Alliaceae, Apiaceae, Asteraceae,

Brassicaceae, Chenopodiaceae, Cucurbitaceae, Fabaceae, Malvaceae, Poaceae, and

Solanaceae) were grown in the greenhouse (35/25 + 5°C day/night temperature, 85 + 5%

RH, with 400 nE/s/m^ light level at midday). These plants were tested for their reaction to

Bipolaris sacchari formulated with an oil-emulsion carrier (Table 4-1). Seeds of the various plant species were either sown directly into plastic pots (10 cm diameter; 8.8 cm height) or

germinated in plastic trays (52.7 cm x 27.5 cm x 6.3 cm) and later transplanted into the

plastic pots. The seedlings were grown in a conmiercial potting medium (MetroMix 300;

Scott's Sierra Horticultural Products Co., Marysville, OH) and were watered regularly as

needed. Seedlings were fertilized with Multicote (15-15-15 N-P-K fertilizer; Scott's Sierra

Horticultural Products Co., Marysville, OH).

Inoculum Production

Spores of B. sacchari were produced by culturing the pathogen on grains of rye

(Secale cereale L.). Rye grains were boiled for 10 minutes and dispensed into glass bottles

(5 cm diameter; 19.5 cm height). The grains were autoclaved for 40 minutes (121°C; 15

psi) for two consecutive days and aseptically inoculated with agar blocks from sporulating 101

12-14 cultures of B. sacchari cultures grown on V-8 agar medium. After incubation for

were harvested by days in the dark at 25 ± TC, black powdery spores that covered the grain rinsing the grain with sterile water. The spore suspension was filtered through two layers of cheesecloth to remove any debris. The spore concentration was determined with the aid of a hemacytometer. A spore concentration of 5 x 10^ spores/ml was used for all inoculations in

the greenhouse. The spores of B. sacchari were formulated with a mixture of 16%

Sunspray® 6E oil (horticultural oil; Sun Company Inc., Philadelphia, PA), 10% light mineral

oil (Fisher Scientific; Fair Lawn, NJ), and 74% sterile water.

An alternative method was employed to harvest spores and store them in a dry form

until inoculation. In this method, a vacuum pump was used to separate the spores from the

water by trapping the spores on filter paper discs or on 10-^m nylon screens. The spores on

the filter paper discs were placed in a petri dish and dried in a 30°C incubator for 3-5 days.

The spores on the nylon screens were dried by hanging the screens inside a lighted fume

hood with a temperature of 27°C for 2 days. After drying, the spores were stored in vials at

10°C. The inoculum for inoculations in the greenhouse was prepared by resuspending the

dry spores in sterile water.

Inoculation

The greenhouse tests were done in the winter of 1999 through the summer of 2000.

All experiments were done twice. Depending on the plant species, inoculations at 3-5 weeks

after seed germination were done with three replications (three pots/species) per treatment,

with each pot containing 3-5 plants, depending on the plant species being tested. Plants 102

were sprayed until full coverage of the foliage was achieved, with hand-pumped sprayers.

Control plants were sprayed with the oil emulsion.

Disease Rating

Inoculated plants were observed for symptoms of disease and phytotoxic damage every week for four consecutive weeks. The rating system used was: 1= immune (none or few specks observed); R= resistant (small lesions and no lesion expansion after 28 days, minor phytotoxic damage from the oils on leaf edges); MS= moderately susceptible (larger lesions or leaf bum observed but recovery occurred within 28 d); S= susceptible (large areas of the foliage burned or blighted but death was not observed within 28 d); HS= highly susceptible (severe blighting of all leaves, plant death observed within 28 d).

Results

Of the 45 plant species tested, 19 species that belonged to six families (AUiaceae,

Apiaceae, Asteraceae, Cucurbitaceae, Poaceae, and Solanaceae) were found to be

moderately susceptible, susceptible, or highly susceptible to the fungus-oil emulsion mixture

(Table 4-1). These susceptible grass and crop species were: Cynodon dactylon

(bermudagrass), Dactyloctenium aegyptium (crowfootgrass), Eleusine indica (goosegrass),

Digitaria sanguinalis (large crabgrass), Pennisetum glaucum (pearl millet), Avena sativa

(oats), Secale cereale (rye), Lolium multiflorum (ryegrass), Cenchrus echinatus (southern

sandbur), Zea mays (sweet com), Paspalum urvillei (vaseygrass), and Triticum vulgare

(wheat), Daucus carota (carrot). Solarium melongena (eggplant). Allium cepa (green onion),

and Lycopersicon esculentum (tomato). Plant species that were moderately susceptible were 103

Cichorium endivia (endive), Cucumis melo (cantaloupe), and johnsongrass {Sorghum halepense) (Table 4-1).

The highly susceptible grass species exhibited severe blighting or plant death, and none of the plants recovered. The susceptible grass species exhibited various degrees of blight but were able to recover from the damage within 28 d. Carrot and green onion sprayed at 15 d after germination exhibited severe foliar bum (first trial) but when sprayed at

28 d after germination (second trial), they were only moderately burned. The damage to carrot and green onion was more severe when the plants were inoculated at a younger stage

(15 d after germination). The damage to eggplant was more severe in the first trial than in the second trial. The reaction of eggplant was characterized as susceptible (leaf bum and some necrotic lesions) to moderately susceptible (lesser leaf area bumed) for the first and

second trial, respectively. Leaves of tomato plants that were inoculated at 14 d (first trial) or

28 d after germination (second trial) were oil-soaked and mostly bumed.

In general, the most severe reactions were observed on a number of graminaceous

plants. Few crop species from Alliaceae, Apiaceae and Solanaceae were either moderately

susceptible or susceptible. All other species exhibited no symptoms or a range of minor

symptoms (specks, small lesions, discoloration from the oil, and leaf-edge bum) that did not

worsen with time. Recovery from the inoculation and growth of new healthy leaves occurred

within 28 d (Figs. 5-1).

Discussion

Bipolaris sacchari is known to cause leaf spots on several species of grasses (Alfieri

et al., 1994; Farr et al., 1989). Hence, the susceptible reaction of several grassy species was 104

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not unexpected. Grasses sprayed with B. sacchari formulated in the oil emulsion did not show typical leaf spots. Instead, they all exhibited severe blighting or death within 28 d.

The combined effects of the pathogen and the phytotoxic effects of the oil emulsion contributed to the severity of the damage.

Formulation of chemical herbicides with oils (as adjuvants), which can affect leaf cuticle, has been used to enhance herbicidal effects (Baker, 1970; McWhorter and

Barrentine, 1988). In the case of bioherbicides, the cuticle damage caused by oils may

facilitate fungal penetration, and result in enhanced virulence of the fungal pathogens and the possible alteration of host range. Spores of Altemaria crassa (Sacc.) Rands, a biocontrol agent for jimsonweed {Datura stramonium L.), when formulated with aqueous

filtrates of jimsonweed or hemp sesbania (Sesbania exaltata [Raf.] Cory) or fruit pectin, were able to infect several crops and weeds that were previously non-susceptible to A. crassa (Boyette and Abbas, 1994).

When the spores were formulated with an invert emulsion (Amsellem et al.,

1991), it abolished the host selectivity of Altemaria cassiae Jurair & Khan, a potential biological control agent for sicklepod {Senna [=Cassid\ obtusifolia [L.] Irwin & Bamaby) in a soybean crop. In host-range tests, spores of A. cassiae suspended in water did not infect soybean {Glycine max [L.] Merr.), peanut {Arachis hypogaea L.), cotton

{Gossypium hirsutum L.), com and 26 other crop and weed species in nine families

(Walker, 1982). Spores of A. cassiae in invert emulsion were able to infect soybean; it

caused lesions with necrotic centers (Amsellem et al., 1991). The invert emulsion also increased the virulence of two avirulent fungi, Trichoderma harzianum Rifai and

Aspergillus nidulans; it allowed them to infect a variety of plants (Amsellem et al., 1991). 109

These results were attributed to the physical damage caused by the invert emulsion to the plant cuticle or the possible suppression of the host defense responses. It is possible that the application of B. sacchari, a pathogen associated with grasses, with oil emulsion has

extended its host range to members outside the Poaceae family.

Aside from the formulation, the age and the vigor of the test plants can also affect their reaction to the fungus-oil emulsion mixture. Plant age has profound effects on disease susceptibility or disease severity (Populer, 1978) and plants grown in greenhouses are often more susceptible to infection than field-grown plants (TeBeest, 1985;

Templeton et al., 1984). Host range tests in the greenhouse are conducted under optimum environmental conditions for infection and disease development, conditions that are not always likely to exist in the field (Watson, 1985). Hence, the host range of a biological

control agent is often broader in studies in the studies in the greenhouse than would occur in the field (Dinoor and Eshed, 1987; Watson, 1985; Zwolfer and Harris, 1971). In a host-range study of Exserohilum monoceras (Drechs.) K.J. Leonard & E.G. Suggs, a potential bioherbicide for Echinochloa species, infections on com occurred only when the inoculated com plants were exposed to 24 h of dew. Under field conditions, com was not infected by E. monoceras (Zhang and Watson, 1997).

Host-range studies done by Chandramohan in 1999 indicated that D. gigantea was pathogenic to some weedy grasses. The fungus, however, was not pathogenic to any of the economically important crop species tested (Chandramohan, 1999).

The use of B. sacchari and D. gigantea as bioherbicides of cogongrass are not

likely to pose risks to nontarget plants if they are used to manage monotypic stands of cogongrass in noncrop areas. Furthermore, since the oil formulation improves the 110

adhesion of the spores to the leaf surface, spores of both isolates are not likely to be dispersed by the wind and carried to nontarget plant species. The disease caused by B. sacchari or D. gigantea on cogongrass does not go into secondary cycles, and therefore these fungi do not produce secondary inoculum that can reach nontarget plants. CHAPTER 6 FIELD EFFICACY OF Bipolaris sacchari and Drechslera gigantea AS BIOHERBICIDES OF COGONGRASS

Introduction

Determination of field efficacy is one of the most important steps in the

evaluation of prospective weed biocontrol agents. Organisms that perform effectively

under controlled environments in the greenhouse or growth chambers may exhibit

variable activity against the target weed under natural conditions in the field (Rosskopf et

al., 1999). Aside from technological difficulties in mass production and marketing of a

commercially acceptable formulation, competition from chemical herbicides and

unprofitable markets, the level of efficacy under field conditions is one of the major

considerations that determine whether a bioherbicide can be registered and used

commercially (Charudattan, 1991).

Numerous weed biocontrol agents have been field tested to determine their

commercial feasibility. Under field conditions, DeVine® {Phytophthora palmivora

[Butler] Butler) was found to control more than 90% of the infestations of stranglervine

{Morrenia odorata [Hook. & Am.] Lindl.) in citrus groves (Ridings, 1986). COLLEGO®

(Colletotrichum gloeosporioides [Penz.] Penz. & Sacc. f. sp. aeschynomene) was shown to provide greater than 85% control of northern jointvetch consistently {Aeschynomene virginica [L.] B. S. P.) in rice and soybean crops. Many other candidate biocontrol

agents have been tested for their field performance (Charudattan, 2001). These

111 112

biocontrol agents include Colletotrichum coccodes (Walk.) Hughes for Abutilon

theophrasti Medic. (DiTommaso et al., 1996), Phomopsis convolvulus Ormeno for

Convolvulus arvensis L. (Ormeno-Nunez, 1988), Phomopsis amaranthicola Rosskopf,

Charud. & DeVal. (Rosskopf, 1998), Altemaria altemata (Fries:Fries) von Keissler for

Amaranthus spp., Drechslera avenacea (M. A. Curtis ex Cooke) Shoemaker ior Avena

fatua L.(Vurro et al., 1999), Stagonospora convolvuli Deam. & House for Calystegia

sepium (L.) R.Br. (Guntli et a., 1999) and Convolvulus arvensis L. (Pfirter et al., 1997),

Phomopsis cirsii Grove on Cirsium arvense (L.) Scop. (Leth and Andreasen, 1999),

Dactylaria higginsii (E. S. Lutrell) M. B. Ellis for Cyperus rotundus L. (Dinoor et al.,

1999; Kadir and Charudattan, 2000), Exserohilum fusiforme Alcorn, E. monoceras

(Drechsler) Leonard et Suggs, and Colletotrichum graminicola (Cesati) Wilson for

Echinochloa spp. (Van Tuat et al., 1998; Yang et al., 2000; Zhang and Watson, 1997),

Cercospora piaropi Tharp. (Vincent and Charudattan, 1999) and Altemaria eichhomiae

Nag Raj & Ponnappa (Shabana et al., 1999) on Eichhomia crassipes (Mart.) Solms ,

Colletotrichum truncatum (Schwein) on Sesbania exaltata (Rafin.) Cory (Boyette et al.,

1999), Fusarium nygamai Burgess & Trimboli (Kroschel et al., 1999), F. oxysporum

SchlechtendahliFries and F. semitectum (Berkeley & Ravenel) var. majus Wollenweber

(Ciotola et al., 1995) for Striga hermonthica (Del.) Benth.

The most frequently cited reason for variable field performance of bioherbicides

is the constraint imposed by the environment after inoculation (Greaves et al., 1998).

Humidity and duration of dew period are the main limiting environmental factors to the

initiation and progress of disease in the field (Auld et al., 1990; McRae and Auld, 1988).

Several bioherbicides have been reported to require more than 12 h of dew to cause high 113

levels of disease (Boyette and Walker, 1985; Makowski, 1993; McRae and Auld, 1988;

Morin et al., 1990; Wymore et al., 1988). Requirements for long periods of moisture may limit the potential use of these bioherbicides in the field, especially in areas with infrequent rainfall and low relative humidity.

The problem with dew requirement has been addressed by formulation of biocontrol agents with moisture-retaining materials or by the timing of application.

Humidity levels may be increased with more efficient wetting agents or invert emulsions

or when the biocontrol agent is applied as a gel or as a granular formulation (Walker,

1981; Walker and Connick, 1983). The application of the mycoherbicide may be timed to occur when the environmental conditions are most favorable for infection. The timing

of sprays to take advantage of rain, dew, and irrigation is one way to overcome the

problem of delay in the onset of dew (Auld et al., 1990; McRae and Auld, 1988;

Makowski, 1993).

Spray volume and spray distribution are also critically important for the efficacy of a bioherbicide. Typically, the plants are sprayed until the excess liquid runs-off to ensure good coverage with the inoculum. This practice optimizes the chances to produce

successful infections but it seriously impedes the assessment of the real potential of the

agent to perform under field conditions. High volumes do not necessarily ensure persistence of the water or the even distribution of the inoculum. Much of the applied

water can run-off the leaf surface, move the spores to the edges of the leaf and sometimes

remove the spores from the infection court (Greaves et al., 1998). Klein and Auld (1995)

showed that the application of Colletotrichum orbiculare (Berk. & Mont.) Arx., a

biocontrol agent for Bathurst burr {Xanthium spinosum L.), at the rate of 1000 1/ha was 114

generally not superior to 500 and 250 1/ha. They concluded that lower spore doses and carrier volumes were adequate where environmental conditions were favorable for disease development. Low rates of application that can cause severe or lethal damage to the target weed are desired to minimize production and application costs of the bioherbicide (Baker and Henis, 1990).

Biocontrol agents have been formulated with oil to improve application efficiency. Oil-based formulations are more viscous than aqueous formulations and they may be more effective to enable the inoculum to stick to the hydrophobic plant surfaces

(Greaves et al., 1998). In a study by Chandramohan (1999), the application of fungal inoculum formulated with an oil emulsion resulted in the highest level of weed control in the field. The high level of biocontrol efficacy in the field was attributed to better coverage of the target weed surfaces, predisposition of the target weed to the pathogen, and the improved diffusion of toxic metabolites that may have been produced by the fungal inoculum.

Another factor that may affect the efficacy of a bioherbicide is the type of application system. In preliminary work by Chappie and Bateman (1997), there were substantial differences in the final amount of deposited droplets that contained the active

ingredient (i.e. spores), the amount of deposited droplets with no active ingredient, and the droplet size when different application systems were used. In their study, they used three application systems (hydraulic flat fan, air blast sprayer, and spinning disc) to apply spore suspensions of the biofungicide Ampelomyces quisqualis Cesati ex Schlechtendahl at the rate of 1.25 x 10^' spores/ha. With the hydraulic flat fan, the percentage of droplets

that contained more than 1 spore was 18.9% while the percentage of droplets without 115

spores was 67.3% (average for three different nozzles). When the air blast sprayer was used, the percentage of droplets with more than 1 spore was 0.5%, while the percentage of droplets without spores was 95.6%. With the spinning disc, the percentage of droplets

with more than 1 spore was 81.5% and 36.5% for the oil- and the water-based formulation. The percentage of droplets without spores was 6.5% and 39.9% for the oil- and water-based formulation. The percentage of droplets greater than 300 ^m in size from the hydraulic flat fan nozzle was 30% for nozzle 2080-14, 50.3% for nozzle 2080-

16, and 76.2 for nozzle 2080-30%. Both the air blast sprayer and the spinning disc did not produce droplets bigger than 300 ^m (Chappie and Bateman, 1997).

The size and the size range of droplets produced by the sprayer are affected by the flow rate of the suspension through the sprayer. Row rate of the suspension through a sprayer depends on the size of the inlet of the nozzle, on whether the suspension is pumped into the atomizer, and on the viscosity of the suspension (Jones, 1998).

Substantial differences may exist between application systems that are used in the laboratory, greenhouse, and the field. Typically, pressurized canisters and hand-pumped sprayers are used in the laboratory or greenhouse as opposed to large spray tanks with pumps and recycling systems or rotary atomisers such as the ultra low volume application

(ULVA) systems that are used in the field. These differences may have significant effects on the eventual distribution of the biocontrol agent through the canopy and significantly change the efficacy of the bioherbicide (Chappie and Bateman, 1997).

This study was done to 1) determine the influence of spore concentration, carrier concentration, and bioherbicide application volume on the field efficacy of B. sacchari and D. gigantea, 2) determine the optimum level of spore concentration, carrier 116

concentration, and application volume for the control of cogongrass under field conditions, 3) determine the effect of the applicator system and application volume on the field efficacy of the bioherbicide agents, and 4) compare the field efficacy of two application volumes of B. sacchari, D. gigantea, and the mixture of the two isolates with the ultra low volume application (ULVA) system.

Materials and Methods

Inoculum Production

Spores of B. sacchari and D. gigantea were obtained from cultures grown on rye grains in glass bottles (5 cm diameter; 19.5 height). Fifty grams of rye grain were inoculated and the cultures were incubated at room temperature (25 + 2 °C) for 12-14 days. The grain-cultures were rinsed with sterile water and the spores were dislodged by vigorous stirring with a clean metal spatula. Spore suspensions were filtered through two layers of cheesecloth to remove debris and the spore concentration was determined with the aid of a hemacytometer.

Freshly harvested spores were either used in the field trials or dried and then stored for future use. Spores that were not used for inmiediate inoculation were collected

by filtration of the spore suspension through a 10-|im nylon screen or a Whatman filter paper no.l (5-cm diameter). Spores on the nylon screens were dried inside a lighted fumehood for 2-3 days (27°C). Spores on the Whatman filter papers were kept inside plastic petri dishes (15 x 150 mm) and incubated for 2-3 days inside a 30°C incubator.

Dry spores were transferred to sterile glass vials or plastic bottles and stored at 10°C.

Inoculum was prepared by resuspending the dry spores in sterile water. 117

The bioherbicide was prepared by mixing equal volumes of spores of B. sacchari or D. gigantea suspended in water and an oil emulsion carrier composed of Sunspray 6E

horticultural oil, Fisher light mineral oil, and water at different proportions of each component, as explained below. Spore suspensions that contained twice the desired amount of spores and oil emulsion that contained double the oil concentration were mixed together to achieve the final formulation with the desired spore concentration and carrier concentration.

1999 Mini-plot Experiments

Four field experiments were established in 1999. Experiments with D. gigantea were conducted at Waldo, FL in April-May and at the Biven's Arm experimental site in

Gainesville, FL in July-August. Experiments with B. sacchari were established at two separate stands of cogongrass at the Biven's Arm site in Gainesville, FL in July-August and October-November. Experiments were conducted in stands of cogongrass that were approximately 0.6 m in height. The area was marked with the use of a 0.25 m frame made of polyvinyl chloride pipes. The comers of the each plot were marked with pin flags. The plots had buffer zones on all sides of at least 0.5 m. Experiments were in a completely randomized design (CRD) with four replications per treatment or in a randomized complete block design with four replications per treatment.

For the experiment at Waldo, the following were tested: two spore concentrations of D. gigantea: 4 x 10^ spores/ml and 2 x 10^ spores/ml; two carrier concentrations: 18% oil emulsion (8% Sunspray® 6E and 10% mineral oil) and 26% oil emulsion (16%

Sunspray® 6E and 10% mineral oil); and two application volumes: 100 ml/0.25 m^ and 118

200 ml/0.25 m^. The treatments for experiments at Gainesville site were similar to the experiment at Waldo but the inoculum concentrations of D. gigantea for the Gainesville

site experiments were 4 x 10^ spores/ml and 8 x 10^ spores/ml. Plots at the Gainesville

site, which were under full sunlight, were inoculated in the late afternoon. At the Waldo

site, which was under the shade, inoculation was done in the morning.

In the first experiment with B. sacchari at the Gainesville site, the following were

tested: two spore concentrations: 5 x 10^ spores/ml and 1 x 10^ spores/ml; two carrier concentrations: 18% oil emulsion and 26% oil emulsion; and two application volumes:

100 ml/0.25 m^ and 200 ml/0.25 m^. The trials also included non-inoculated control plots and plots treated with oil emulsion only (18% and 26% oil). The second trial was similar

to the first trial in all details except the spore concentrations which were tested at 5 x

10^/ml and 1.5 x 10^/ml. Inoculations were done with the use of hand-pumped sprayers, in late afternoons.

2000 Field Trials

The following experiments were done at the Biven's Arm experimental site in

Gainesville, FL from September to December 2000. The cogongrass stand was

approximately 1 .3 m in height. Plots were laid out with the use of a m^ wooden frame and the comers were marked with pin flags. The buffer zone between plots was at least

0.5 m. The experiments were carried out in a completely randomized design with four replications per treatment. All inoculations were done in late afternoons. 119

Effect of Applicator System and Application Volume on the Field Efficacy of the Bioherbicide

Two applicator systems, the ultra low volume application system (Micron

ULVA+; Micron Sprayers Limited, Bromyard Industrial Estate, Bromyard, Herefordshire

HR7 4HS, UK) (ULVA) and an air sprayer (Model L-300 Electric Platform Sprayer;

Weed Systems Inc., Keystone Heights, FL) and two application volumes (100 ml/m^ and

200 ml/m^) were tested in this experiment. The test isolate used, B. sacchari, was

formulated with an oil-emulsion carrier that contained 26% oil (16% Sunspray 6E and

10% mineral oil). The spore concentration was 1.5 x 10^ spores/ml.

Comparison of the Field Efficacy of B. sacchari, D. gigantea and B. sacchari + D. gigantea Mixture using the Ultra Low Volume Applicator System

Based on the results of the previous experiment, another trial was performed to

compare two application volumes of both isolates, applied singly or as a mixture, with the

ULVA system. Spores of B. sacchari and D. gigantea (9 x 10^ spores/ml) were

formulated with an oil emulsion that contained 26% oil and applied at the rate of 200

ml/m^ and 300 ml/m^.

Data Collection and Disease Assessment

The inoculated and control plants were observed for symptoms of disease and

phytotoxicity damage at one week after inoculation (WAI) and disease severity or

phytotoxicity damage (DS) was assessed at 2 WAI when the disease symptoms were

more pronounced. Disease severity was visually estimated based on the percentage of

blighted aboveground biomass per plot (0-100 %). The visual estimates were classified 120

into classes using the Barratt-Horsfall scale (Barratt and Horsfall, 1945). The classes in the Horsfall-Barrat-scale correspond to different levels of disease severity. These classes

are: 0 = 0; 1 = 0-3%; 2 = 3-6%; 3 = 6-12%; 4 = 12-25%; 5 = 25-50%; 6 = 50-75%; 7 =

75-88%; 8 = 88-94%; 9 = 94-97%; 10 = 97-100% and 1 1 = 100%. The mean class value was used to determine the disease severity value. Disease assessments were done every week from the 2"'' WAI until the 6'*' WAI. Data were collected similarly for all mini-plot

trials and all other succeeding field trials.

Data Analysis

Data from all mini-plot trials were analyzed using the DS ratings that have been transformed using the Barratt-Horsfall scale. Analysis of variance (ANOVA) with the

General Linear Models (SAS Institute, Gary, NG) and the Mixed Procedures were used to determine the effect of each of the factors an their interactions. Means were compared using the Least Significance Difference test (LSD) with P=0.05.

Results

Influence of Spore Concentration, Emulsion Concentration, and Bioherbicide Application Rate on the Field Efficacy of Drechslera gigantea

Data from the experiments in April and July were analyzed separately because of

significant trial effects. For both trials, there was a significant interaction effect of the

application volume, oil concentration, and the spore concentration (Tables 6-1 and 6-2).

In the trial at Waldo, the highest DS levels (>70%) resulted in treatments with 200 ml of

the following mixtures: 5 x lO' spores/ml and 2x10^ spores/ml + 26% emulsion 121

Table 6-1. Analysis of variance for disease severity of cogongrass (Waldo site) biiect Mum Tip 1 V CllUW' Pr > F 3 98'^0 Block J Ito 1 98 0

1 48 40 69 <--0 0001 Volume 1 I'to 1 148 <-0 Oil concentration 1 9Q 9S 0001

1 48 14^ 86 <-'n 0001 Spore concentration z 1 1J.ou A 148 60 98

148 1 "^4 0 9000 Block X time after inoculation IZ Ito 1 . Jt 1 volume X on conceniraiion i 1481 to 9 67 0 1044 Volume X spore concentration L 148ito Zt.94 J J

Volume X oil concentration x spore 2 148 10.17 <0.0001 concentration Volume X oil concentration x spore 28 148 0.74 0.8236 concentration x time after inoculation Num DP = numerator degrees of freedom. Den DP = denominator degrees of freedom. 122

Table 6-2. Analysis of variance for disease severity of cogongrass inoculated with D. gigantea (Gainesville site) Effect NumDF DenDF F value Pr>F

VolumeT \J 1 U 1 ft 1 36 151.13 <0.0001 90.61 <0.0001 Oilft ft concentrationft l^^^^ft 1 Lft %h4Va V^ft ft 1 36

\JSnnreL/v/ft WV/lftconcentrationvWftftkft Ul-ftV.'ftA 2 36 24.99 <0.0001 Time after inoculation 4 168 1.61 0.1728

\v/ftft LA 4Akft ft ft 44.59 <0.0001 VolumeT V/ft kftft ft ft^^ X oil concentrationLm\^^^%.l 1 36 Volume X spore concentration 2 36 30.74 <0.0001 Volume X time after inoculation 4 168 0.22 0.9277 Oil concentration x spore concentration 2 36 28.96 <0.0001 Oil concentration x time after inoculation 4 168 0.20 0.9391 Spore concentration x time after 8 168 1.07 0.3850 inoculation Volume X oil concentration x spore 2 36 9.43 <0.0005 concentration Num DF = numerator degrees of freedom. Den DF = denominator degrees of freedom. 123

A

18(100) 26(100) 18(200) 26(200)

Oil concentration (%) and appiication volume (ml)

B

80 T 70 4x10 t: 60 8x10 S 50 1 40 S 30 CO 8 20 o 10 ImJllL 18(100) 26(100) 18(200) 26(200)

Oil concentration (%) and application volume (ml)

Fig. 6-1. The interaction effect of spore concentration, emulsion concentration, and application volume on the field efficacy of Drechslera gigantea. Graphs show the results from the April 1999 trial in Waldo, FL (A) and the July 1999 trial in Gainesville, FL (B). 124

(71% and 91.2%). Greater than 70% DS was also obtained with 2 x 10^ spores/ml applied with 26% emulsion at 100 ml/0.25 m^ (Fig. 6-1 A). Time had a significant effect on the level of disease in the field (Table 6-1). The average level of disease caused by all the treatments was higher at 4 to 6 WAI than at the beginning of the experiment (data not shown).

In the trial at Gainesville, the highest DS level (77.8%) was achieved only when

200 ml of 8 X 10^ spores/ml + 26% emulsion was applied. Application of the other spore and emulsion mixtures and the emulsion alone at either rates (100 ml/plot or 200 ml/plot) caused lower levels of foliar blighting (<50%) (Fig. 6-1 B). Time did not have any significant influence on the level of disease in this trial (Table 6-2).

Influence of Spore Concentration, Emulsion Concentration, and Bioherbicide Application Rate on the Field EfHcacy of Bipolaris sacchari

'a 4. Data from the two trials with B. sacchari were analyzed separately due to

significant trial effects. In the first trial, there was a significant interaction effect of the volume and the spore concentration and a significant interaction effect of the oil and spore concentrations (Table 6-3). The highest DS levels (84-86%) resulted from

treatments with 200 ml of the oil emulsion carrier containing 5 x 10^ or 1 x 10^ spores/ml

(Fig. 6-2 A). When the application volume was 100 ml, the application of 5 x 10^ or 1 x

lO^spores/ml spore in the emulsion mixtures resulted in significantly lower DS levels

(47-50%). Plots treated with the emulsion alone (no spores) exhibited <10% foliar

blighting. The DS level caused by the application of 1 x 10^ spores/ml with the higher emulsion concentration (26%) was not significantly higher than the DS level caused by the application of 5 x 10^ spores/ml + 18% or 26% emulsion (Fig. 6-2 B). The effect of 125

Table 6-3. Analysis of variance for disease severity of cogongrass inoculated with B. sacchari (Gainesville site trial 1) Effect NumDF DenDF F value Pr>F

\7V VJlUlli&r^\\^rx\p^ 1 35 186.66 <0.0001

r*f\T\ot^T\\TC\\\c\X\ 1 \J\.iOil CUllL-CllLl allUll X 35 5.79 0.0215 ^nnrp pnnrpntrfltinn 2 35 501.95 <0.0001

Ximp'1 line all&laftpr lllUdliaiiv/iiinnpnlfltinn 4 140 10.76 <0.0001

Volume x oil concentration 1 35 1.78 0.1904

\/r^liim#>V UlUlllC AY olJUlCcnore ^./vllW-tilllponppntrjitionclllUll 2 35 47.44 <0.0001 Vnliimp y time after inoculation 4 140 0.96 0.4338 Oil concentration x spore concentration 2 35 6.77 0.0033 Oil concentration x time after inoculation 4 140 0.02 0.9994 Spore concentration x time after 8 140 1.21 0.2953 inoculation Volume x oil concentration x spore 2 35 1.81 0.1787 concentration Volume X oil concentration x spore 28 140 0.50 0.9838 concentration x time after inoculation Num DF = numerator degrees of freedom. Den DF = denominator degrees of freedom. 126

Table 6-4. Analysis of variance for disease severity of cogongrass inoculated with B. sacchari (Gainesville site trial 2) Fffert Num DF Den DF F value Pr>F V oiumc 1 36 173.20 <0 0001 cuncciiiraiiuii 1 36 7.05 0.0117 7 oporc Luncciiiraiiuii 36 667\J\J 1 56 <0 0001

1 line ciiLCi liiijc-uiciiiv^ii 4 144 14.51 <0 0001 VUIUIIIC A Ull CUIlCCUllallUIl 1X 36 5 72 0.0221

\/r»liitTif* Y cnnrp ponppntrfltinn 2 36 28.49 <0vy « 0001vy V.^ i A volume A iinic diicr iiiuL'Uiaiiuii 144 0 74 0 5643

\J\i CUIlCCIlirallUll A apUlC CUllCCIlllaliUU 36 1 31 0 2814 Oil concentration x time after inoculation 4 144 0.18 0.9509 Spore concentration x time after 8 144 1.54 0.1482 inoculation Volume X oil concentration x spore 2 36 3.60 0.0376 concentration Volume x oil concentration x spore 28 144 0.46 0.9914 concentration x time after inoculation Num DF = numerator DF. Den DF = denominator DF. 127

A

Number of spores^ml

B

Number of spores/ml

Fig. 6-2. The interaction effect of the application volume and emulsion concentration (A) and emulsion concentration and spore concentration (B) on the field efficacy of Bipolaris sacchari. 18(100) 26(100) 18(200) 26(200)

Oil concentration (%) and application volume (ml)

Fig. 6-3. The interaction effect of spore concentration, emulsion concentration, and application volume on the field efficacy of Bipolaris sacchari. 129

1 WKBMIM^'^^

1 ^BB^B^K^^''fiM^Mlt'ii ii f ft

Fig. 6-4. Effect of the interaction of spore concentration, emulsion concentration, and the application volume on the biocontrol efficacy of Bipolaris sacchari on cogongrass in the field. Top row: plot treated with 100 ml of 10^ spores/ml + 18% oil (left); plot treated with 200 ml of 10^ spores/ml + 18% oil (middle); and plot treated with 100 ml of 10^ spores/ml + 18% oil (right). Middle row: plot treated with 200 ml of 10^ spores/ml + 18% oil (left); plot treated with 100 ml of 10^ spores/ml + 26% oil (middle); and plot treated with 200 ml of 10^ spores/ml + 26% oil (right). Bottom row: plot treated with 100 ml of 10^ spores/ml + 26% oil (left); and plot treated with 26% oil only. Photos were taken at the Biven's Arm site in Gainesville, at 12 days after inoculation. 130

the interaction of spore concentration, emulsion concentration, and the application volume on the efficacy of B. sacchari is shown on Fig. 6-4.

In the second trial, interaction of the application volume, oil concentration, and spore concentration had a significant effect on field efficacy of B. sacchari (Table 6-4).

Higher DS levels (70-94%) were achieved with the application of 100 ml of the spore and oil emulsion mixture that contained 1.5 x 10^ spores/ml and 26% oil and 200 ml of the spore and oil emulsion mixture that contained 18% or 26% oil and 5 x 10^ or 1.5 x

10^ spores/ml (data not shown). Application of the spore and emulsion mixtures at the lower rate (100 ml) caused significantly lower DS levels (Fig. 6-3). Plots sprayed with the carriers alone (100 or 200 ml) had the lowest level (<20%) of damage.

In both trials, time had a significant effect on the level of disease (Tables 6-3 and

6-4). In the first trial, DS levels were decreased after 3 WAI. However, in the second

trial, there were higher DS levels from 3 to 6 WAI. Since the trials were done on the same location and the two separate cogongrass stands were not morphologically different, the differences in the results were attributed to the time of inoculation and the differences in the levels of spore concentrations and the quality of the spores used.

Effect of Applicator System and Application Volume on the Field EfRcacy of Bipolaris sacchari

The field efficacy of B. sacchari was significantly influenced by the interaction of the type of applicator system and the application volume (Table 6-5). The highest level of disease (>50%) was achieved with the application of 200 ml of the B. sacchari and emulsion mixture using the ULVA (Fig. 6-5). Application volumes of 100 ml with the 131

air-assisted sprayer or the ULVA and 200 ml with the air-assisted sprayer caused significantly lower DS levels (<35%) (Fig 6- 6). The disease severity was also

Table 6-5. Analysis of variance for disease severity of cogongrass inoculated with B. sacchari Effect Num DF DenDF F value Pr>F

Applicator system 1 60 5.40 0.0236

Application volume 1 60 22.00 <0.0001 Time after inoculation 4 60 9.66 <0.0001

Applicator system x volume 1 60 8.63 0.0047 Applicator system x time after inoculation 4 60 0.03 0.9983 Applicator system x volume x time after 8 60 0.13 0.9979 inoculation Num DF = numerator degrees of freedom. Den DF = denominator degrees of freedom. 132

Fig. 6-5. The effect of applicator system (air-assisted sprayer or ultra low volume applicator [ULVA]) and application volume (100 or 200 ml) on the field efficacy of B. sacchari. 133

Table 6-5. Analysis of variance for disease severity of cogongrass inoculated with B. sacchari

Effect Num DF Den DF F value Pr>F Annlication volume 1 16 75.88 <0.0001 Treatment 2 16 30.55 <0.0001 Time after inoculation 4 64 19.39 <0.0001 Application volume x time after 4 64 0.28 0.8929 inoculation Application volume x treatment 2 16 13.82 0.0003 Application volume x treatment x time 16 64 0.76 0.7272 after inoculation Num DF = numerator degrees of freedom. Den DF = denominator degrees of freedom. 134

B. sacchari D. gigantea B. sacchari + D. gigantea

Treatment

Fig. 6-6. The field efficacy of B. sacchari, D. gigantea applied singly or as a mixture at the rate of 200 ml/m^ or 300 ml/m^ using the ultra low volume application (ULVA) system. 135

Fig. 6-7. Field efficacy of Bipolaris sacchari, Drechslera gigantea, and the mixture of the two isolates applied at two rates with the ultra low volume application (ULVA) system. Top: plots treated with 200 ml of B. sacchari (left), D. gigantea (middle), and the mixture (right). Bottom: plots treated with 300 ml of B. sacchari (left), D. gigantea (middle), and the mixture (right). Photos were taken at the Biven's Arm site at Gainesville, FL at 6 weeks after inoculation. 136

influenced by time; in general, there was a downward trend in the level of disease from 2

WAI to 6 WAI (data not shown).

Comparison of the Field Efficacy of B. sacchari, D. gigantea and Mixture of B. sacchari and D. gigantea with the Ultra Low Volume Applicator System

The lowest disease severity (41.63%) resulted from the application of 2(X) ml of

D. gigantea spores formulated with the emulsion (Fig. 6-6). Application of D. gigantea at a higher rate (300 ml/plot) and the application of B. sacchari and the mixture of the two isolates at the rate of 200 ml/plot or 300 ml/plot caused higher DS levels (>70%). At the application rate of 300 ml/plot, the field efficacy of both isolates and their mixture were not significantly different. However, at the rate of 200 ml/plot, B. sacchari and the mixture of B. sacchari and D. gigantea had a higher biocontrol efficacy than D. gigantea.

Disease severity significantly changed over time, but unlike in the previous experiment, a downward trend was not observed, and the disease severity levels were higher (>70%) from 3 to 6 WAI (data not shown). The efficacy of the two isolates and their mixture at the two application rates are shown on Fig. 6-7.

Discussion

Bipolaris sacchari and Drechslera gigantea were able to cause disease on cogongrass under field conditions in repeated trials in 1999 and 2000. Inoculated cogongrass exhibited yellowing and brown foliar lesions within five days after inoculation. Distinct blighting of the leaves was observed at 2 WAI. In general, higher disease severities were achieved in these mini-plot trials when a higher volume of the spore and emulsion mixtures was used. When a higher volume was applied, even 137

mixtures with a lower spore concentration (10^ spores/ml) and a lower emulsion concentration (18% oil) caused high levels of disease on cogongrass.

Decrease in DS levels (expressed as percentage of blighted biomass) over time resulted from the appearance of new green and healthy leaves or shoots. Generally, disease severities were significantly reduced between 3 to 6 WAI, except in experiments done in April -May 1999 and October -November 1999 and 2000, when the disease levels followed an upward trend, with disease severity significantly increased between 3 to 6 WAI.

Two factors that might have contributed to this upward trend of disease severity are the temperature and the amount of rainfall during the experiment. Experiments where the disease severity increased with time had similar temperature regimes, the maximum temperature ranged from 26-27.5°C and the minimum temperature ranged from 10-12°C, in both experiments. The amount of total rainfall ranged from <1.0 inch to 2.25 inches.

In controlled environment studies, dry weight of cogongrass, plant height, and leaf area increased when the temperature regime was 30/25°C as compared to other temperature

regimes (24/1 8°C or 27/22°C) (Wilcut et al., 1988). Cooler temperatures and water stress may have slowed down the production of new healthy leaves.

In one of the experiments that showed a downward trend in disease severity over time, the maximum and minimum temperatures were 34.7°C and 22.3°C and the total rainfall was 9.05 inches. However, in the other experiment with the same downward

trend, the maximum and minimum temperatures were 28.8°C and 1 1 .3°C and the rainfall was 4.22 inches. The level of foliar blighting was influenced by factors other than the

differences in spore concentration within and between trials, emulsion concentration, ' 138

application volume, temperature, and rainfall. Other factors that may have contributed to the variability of results include the location of the study, the type of cogongrass population, and the quality of the spores used in each trial.

Complete coverage of the cogongrass foliage with the bioherbicide is necessary to achieve high levels of foliar blighting in the field. Water volumes are important for the uniform application of inoculum (Mortensen and Makowski, 1989). According to Shrum

(1982), proper inoculum distribution sufficiently early or uniformly is essential to create

epidemics. For a weed like cogongrass, which can produce numerous leaves that can

grow as tall as 3 m high, a high application volume is needed to ensure complete and

extensive coverage of the massive foliage. Complete coverage of the leaf surfaces with

inoculum is also important because B. sacchari and D. gigantea do not produce

secondary infections on cogongrass, and therefore there are no additional sources of

inoculum. The level of cogongrass control in the field ultimately relies on the level of

initial infection.

Enhancement of biocontrol efficacy by application of larger volumes of inoculum

has also been demonstrated by Imaizumi et al. in 1997. There was a distinct dose-

response effect of cell concentration of Xanthomonas campestris (Pammel) Dowson pv.

poae (strain JT-P482) and water volume on annual bluegrass (Poa annua L.). At cell

concentrations as low as 10^ CFU/ml, the application volume significantly decreased the

percentage of annual bluegrass control. Acceptable levels of control (>75% at 4 weeks

Q after inoculation) were achieved when 10 CFU/ml inoculum was applied at the rate of

400 ml/m\ lO' CFU/ml inoculum applied at the rate of 100-400 ml/m^, or when 10'°

CFU/ml inoculum applied at the rate of 50-400 ml/m^. The rate of disease caused by X. 139

campestris pv. poae strain on annual bluegrass was accelerated by the application of larger volumes of bacterial cell suspensions. The recommended application rate for the

10'° commercial use of this bioherbicide is of 4 x cells/ m^or 10^ CFU/ml x 400 ml/m^

(Imaizumi et al., 1997).

Klein and Auld (1995) reported that higher water volumes favored disease development by Colletotrichum orbiculare on Bathurst burr {Xanthium spinosum).

However, they also observed that when environmental conditions were conducive to disease development, the lower spore doses and carrier volumes were adequate.

Mortensen and Makowski (1989) observed that the application of higher volume of

water, which was used as the carrier for Collectotrichum gloeosporioides (Penz.) Sacc.

round-leaved mallow {Malva f. sp. malvae, produced a more uniform initial infection on pusilla L.) in the field.

The level of control of cogongrass was also improved by increased coverage of

the foliage of cogongrass with chemical herbicides. In tests in the greenhouse, the shoot

biomass, rhizome biomass, and shoot regrowth of cogongrass decreased when foliar

coverage with glyphosate increased. Increased coverage of cogongrass with imazapyr

also resulted in decreased rhizome biomass (Chase et al., 1996). In field trials, imazapyr

provided greater level of control (from 20% to 40%) when the application rate increased

from 46 1 to 234 1/ ha (Willard et al., 1997). In dense stands of cogongrass, greater

coverage was more important than diluent concentration.

Realistically, low application volumes are preferred because they reduce the

volume and weight to be transported to the target site and the time needed for application.

Higher volumes are aimed to provide complete wetting of the target surface, but this also 140

causes high run-off spray from the target. High application volumes do not always result

in high levels of disease due to low application efficiency. Two main parameters that influence effective application are the application equipment and the formulation of the

inoculum. The type of application equipment and the formulation of the inoculum are

interrelated; the equipment will not perform optimally if the characteristics of the product

are not suitable, or vice versa (Jones and Burges, 1998).

In this study, two application systems were used to apply spores of B. sacchari

formulated in oil emulsion at the two rates 100 ml/m^ and 200 ml/m^. The interaction

between the type of applicator and the application volume influenced the field efficacy of

B. sacchari formulated in 26% oil. The highest level of disease (>50%) was achieved

with the ULVA system and application volume of 200 ml of the spore and oil emulsion

mixture to a m^ plot. When applied with an air sprayer, the same volume of inoculum

caused significantly lower levels of disease (<35%).

The use of ULVA to apply the two promising isolates at lower application

volumes (200 and 300 ml/m^) was further explored. With the ULVA system, B.

sacchari, and a mixture of B. sacchari and D. gigantea that contained 10^ spores/ml

caused >70% disease severity on cogongrass when the application volume was 20O

ml/m^. With a higher application volume (300 ml/m^), the two isolates and their mixture

caused the same level of disease (>70%).

The ULVA may be used to apply lower volumes of B. sacchari or a mixture of B.

sacchari and D. gigantea and still achieve >70% disease severity on cogongrass. In the

mini-plot trial reported here, satisfactory levels of control were achieved when the

application volume was 200 ml/0.25 m^ (800 ml/m^) using hand-pumped sprayers. With 141

the ULVA system, a satisfactory level of control (>70%) was achieved even when the application volume was reduced to 200 ml/m^. The type of application system might circumvent the problem of large volume requirements in the field. CHAPTER 7

SUPPRESSION OF COGONGRASS WITH BIOHERBICIDE APPLICATION AND PLANT COMPETITION

Introduction

An integrated management approach that utilizes several methods of control is needed to manage cogongrass effectively and increase the likelihood of its

suppression. Cogongrass is able to re-invade an ecological niche that is not filled with another plant species after control methods have been implemented. Thus, the establishment of desirable species that will replace cogongrass is essential for long-

term control (Shilling et al., 1997).

Cover crops or tree hedgerows have been used in West Africa and Asia to control cogongrass. The use of cover crops to control cogongrass is based on the sensitivity of cogongrass to shading. Alley cropping with leucaena {Leucaena leucocephala (Lam.) de Wit) and mother of cocoa (Gliricidia sepium (Jacq.) Walp).

reduced cogongrass dry matter by >90%, compared to natural fallow (Anoka et al.,

1991). Since cogongrass tends to flourish in infertile soil, leguminous crops have been used in West Africa and Asia to increase soil nitrogen and to control cogongrass. Legumes used to control cogongrass include calop {Calopogonium mucunoides Desv.), centrosema (Centrosema pubescens Benth.), stylo (Stylosanthes guyanensis [Aubl.] Sw. CV. Cook), tropical kudzu (Pueraria phaseoloides (Roxb.)

142 143

Benth.), sunn hemp (Crotalaria juncea L.), and jackbean {Canavalia ensiformis [L.])

(Anonymous, 1995; Guritno et al., 1992).

Gaffney (1996) used bermudagrass {Cynodon dactylon [L.] Pers.) and hairy indigo {Indigofera hirsuta Harvey) combined with glyphosate or imazapyr to suppress reinvasion by cogongrass Florida, USA. Both plant species grew vigorously and remained free of cogongrass for up to 2 years after seeding. The success of this strategy relies on the tolerance of the desirable species to herbicides and soil type.

Aside from bermudagrass and hairy indigo, bahiagrass has also been considered as a promising species that can be used to suppress cogongrass (Shilling et al., 1995).

A native of eastern Argentina, bahiagrass {Paspalum notatum Fluegge var.

saurae Parodi "Pensacola") has become one of the major forage grasses in the

southeastern United States (Burton, 1967; Quarin et al., 1984). In Florida, bahiagrass

is grown as forage more than any other pasture species, covering approximately 1

million ha (Chambliss, 1996). Bahiagrass is also the most common turf species used

for soil stabilization and beautification on Florida rights-of-ways (Beard, 1980). It is

extensively planted from seed along highways of Florida, North Carolina, and other

subtropical and mild temperate areas (Anonymous, 2000). Bahiagrass is a warm-

season perennial with a deep fibrous root system that can extend to 8 feet deep. It is

well adapted to drier, infertile soils and does best at pH 5.5 to 6.5. Bahiagrass turf

requires little or no irrigation and minimal fertilization, 5 to 10 g of nitrogen per m^

per year, and has few pest problems (Anonymous, 2000). However, it grows poorly

in areas with moderate or heavy shade. Because of their slow growth rate, seedlings

are weak competitors and are susceptible to weed invasion (Busey and Myers, 1979;

Watson and Burson, 1985). Because cogongrass becomes established more rapidly 144

than bahiagrass seedlings, large land areas planted to bahiagrass have been infested or are at risk of being infested with cogongrass (Willard and Shilling, 1990; Shilling et

al., 1997).

In host range studies in the greenhouse, it was determined that bahiagrass was resistant to the biological control agent Bipolaris sacchari. The objective of the

following study is to determine if B. sacchari could be used to suppress cogongrass in mixed plantings of cogongrass and bahiagrass and allow the successful establishment of bahiagrass.

Materials and Methods

Test-Plant Propagation

Cogongrass plants were propagated from rhizomes collected from a cogongrass patch near Lake Alice on the University of Florida campus, Gainesville,

FL. Bahiagrass was grown from seeds obtained from Dr. G. Miller of the Department of Environmental Horticulture, University of Florida, Gainesville, FL. All tests plants were grown in plastic pots (27-cm diameter; 23-cm height) that contained a commercial potting medium (MetroMix 300; Scott's Sierra Horticultural Products

Co., Marysville, OH) and were watered regularly. The plants were grown in a greenhouse at 35/25 + 5°C day/night temperature, 85 + 5% relative humidity and 400

|iE/m^/s natural light intensity at midday.

In the first experiment, mature cogongrass and bahiagrass plants, consisting of

one erect tiller with ca. 12-cm long root system (referred to herein as "planting material") were transplanted in the potting medium and clipped 5 cm above the soil line two days after transplanting to stimulate regrowth. In the subsequent 145

experiments, cogongrass and bahiagrass planting materials were trimmed and weighed before planting in the potting medium. All plants were fertilized with 10 g of a 15-15-15 N-P-K fertilizer (Multicote; Scott's Sierra Horticultural Products Co.,

Marysville, OH).

Inoculum Preparation

Bipolaris sacchari was cultured on sterilized rye grain for 12-14 days at room temperature as described in chapter 3. The inoculated grains were rinsed with sterile

water to recover the spores. The spores were collected by filtering the spore

suspension through a 10-|xm nylon screen (Nytex screen; Sefar America Inc., Depew,

NY) and the screen and spores were air dried in a lighted fumehood without air flow

for 1-2 days. The dry spores were transferred to autoclaved vials and stored at 10°C

until use. Inoculum for this study was prepared by resuspending dry spores of B.

sacchari in sterile water (72%), Sunspray® 6E horticultural oil (16%), and light

mineral oil (10%). The spore concentration was determined with the aid of a

hemacytometer.

Effect of Inoculation ofB. sacchari in an Oil Formulation on Bahiagrass- Cogongrass Mixtures

The bahiagrass-cogongrass mixtures were inoculated in the greenhouse at 12

days after clipping. Three pots planted with the cogongrass-bahiagrass mixture were

sprayed with the emulsion formulation of B. sacchari that contained 7x10^

spores/ml. The inoculum was applied with a hand-pumped sprayer until the foliage

was completely covered. Another set of cogongrass-bahiagrass mixtures was not inoculated and was used as the control treatment. Inoculations were done in late afternoons. Inoculated bahiagrass-cogongrass mixtures were observed for infection at

24 h after inoculation. Thereafter, the plants were observed every seven days for 28 days. The experiments were done, twice, in the fall of 2000.

Effects of Plant Competition and Plant Competition Plus Application of an Oil Emulsion Formulation of B. sacchari on the Growth of Cogongrass and Bahiagrass

December 2000-March 2001 Experiments

Four sets of plants (3 pots/set) were prepared for this experiment. One set was planted with cogongrass only, one set was planted with bahiagrass only, and two sets were planted with equal numbers of cogongrass and bahiagrass planting materials with comparable fresh weights. All inoculations were done in a greenhouse at 35/25

+ 5°C day/night temperature, 85 + 5% relative humidity, and 400 ^lE/m^/s natural light intensity at midday. The plants were watered as needed.

Only one set of the bahiagrass-cogongrass mixtures was inoculated with the

26% oil-emulsion formulation of B. sacchari that contained 5 x lO' spores/ml. All other plants were not inoculated and were used as controls. Non-inoculated controls

were. For both trials, the bahiagrass-cogongrass mixtures were reinoculated at 5 days

after the first inoculation (DAI) with the leftover inoculum to ensure disease development. Test plants were harvested and weighed at 12 weeks after the second inoculation. The effect of plant competition and plant competition plus B. sacchari oil-emulsion formulation on bahiagrass and cogongrass growth was measured in terms of the combined underground and aboveground biomass. The difference

between the final fresh weight (taken at 12 wk after the second inoculation) and the 147

initial weight (taken before planting) was used in the data analysis. The data from two

trials were pooled to get a better estimate of the error term, and analyzed. Means were compared using Duncan's Multiple Range test at P=0.05.

March-May 2001 Experiments

Two sets of plants (6 pots/set) were prepared for this experiment, one set was

planted with cogongrass only, one set was planted with equal numbers of cogongrass

and bahiagrass (3 plants per grass species). Each of the pots were fertilized with 10 g

of Multicote 15-15-15 N-P-K fertilizer and watered regularly. At 5 weeks after

planting, three pots with cogongrass and cogongrass-bahiagrass mixture were

inoculated with spores of B. sacchari (7 x 10^ spores/ml) formulated with an oil

emulsion containing 26% oil. Non-inoculated plants were used as controls. In the

first trial, the plants were inoculated twice to produce a high level of disease. In the

second trial, the plants that have been inoculated once were incubated in a dew

chamber (27°C, 100% RH) for 7 h to ensure successful infection. Test plants were

observed weekly from the first to the 5* week after inoculation (WAI). At the 5*

WAI, cogongrass biomass (aboveground and belowground), plant height, and

rhizome production were determined. The data for two trials were pooled to obtain a

better estimate of the error term and analyzed. Means were compared using Duncan's

Multiple Range test at P=0.05. Bahiagrass biomass in inoculated and non-inoculated

cogongrass-bahiagrass mixtures was also determined. The weights of bahiagrass

from two the trials were pooled and analyzed using the Wilcoxon Rank Sum test at

P=0.05. 148

Results

Effect of Inoculation with B. sacchari in an Oil Emulsion on Cogongrass- Bahiagrass Mixtures

At 3 DAI, inoculated cogongrass exhibited oil-soaking, discrete and

coalescent leaf lesions, and blighting. Bahiagrass did not have any lesions but

exhibited some degree of oil-soaking. At 5 DAI, cogongrass plants were severely

blighted, and bahiagrass exhibited some minute foliar specks but blighting was not

observed. Cogongrass plants were either dead or severely blighted and no healthy

regrowth was observed within 4 weeks after inoculation (Fig. 7-1). Similar results

were obtained from the two trials.

Effects of Plant Competition and of Plant Competition plus Application ofB. sacchari on the Growth of Cogongrass and Bahiagrass

December 2000-March 2001 Experiments

The first inoculation did not produce any disease symptoms or phytotoxic

damage on either plant species after 3 days. Cogongrass exhibited oil-soaking and

lesions only at 3-4 days after the second inoculation. Inoculated cogongrass plants

were severely blighted at 7 days after the second inoculation, while bahiagrass plants

showed some phytotoxic damage. For both trials, the inoculated cogongrass plants

were either severely blighted or dead at 12 weeks after the second inoculation. No

cogongrass regrowth was observed in the first trial. However, in the second trial, a

few new cogongrass plants emerged from the inoculated bahiagrass-cogongrass

mixtures, which meant that some of the rhizomes were not weakened by the

inoculation and were still able to produce new shoots. There were significant 149

Fig. 7-1. Suppression of cogongrass with Bipolaris sacchari and plant competition. Top: cogongrass (arrows) exhibiting severe blighting caused by B. sacchari and healthy bahiagrass. Middle: cogongrass-bahiagrass mixture at 2 weeks after inoculation with B. sacchari (left) and uninoculated mixture of cogongrass and bahiagrass (right). Bottom: cogongrass-bahiagrass mixtures at 12 weeks after inoculation with B. sacchari showing no regrowth of cogongrass (left), and bahiagrass growth ranging from sparse (middle) to dense (right). Table 7-1. Analysis of variance for cogongrass and bahiagrass fresh weight (December 2000- March 2001 experiments) Cogongrass fresh weight Bahiagrass fresh weight

Source Treatment Treatment DF 2 2 SS 264668.44 403958.33 MS 132334.22 201979.17 F value 26.01 43.70 Pr>F <0.0001 <0.0001

DF= degrees of freedom. SS = sum of squares. MS = mean square error. 151

S 450

Cogongrass Bahiagrass Plant species

Fig. 7-2. Effect of plant competition and plant competition plus bioherbicide application on the growth of cogongrass and bahiagrass. Within each plant species, bars with the same letters are not significantly different at P=0.05, as determined by Duncan's Multiple Range test. 152

treatment effects on cogongrass and bahiagrass fresh weights (Table 7-1). Biomass of the inoculated cogongrass plants was significantly lower than the biomass of the non-inoculated cogongrass (Fig. 7-2). Not all of the bahiagrass plants were able to

recover from the phytotoxic damage inflicted by the oil emulsion. The growth of

inoculated bahiagrass planted with cogongrass ranged from sparse to dense (Fig. 7-1).

Without cogongrass and B. sacchari, the average biomass of bahiagrass was

395.67 g/pot (Fig. 7-2). When planted with cogongrass, the biomass of bahiagrass

was significantly lower (1 16.5 g/pot) (P=0.05). However, no further reduction in

biomass was observed when the bahiagrass planted with cogongrass was inoculated

with the oil emulsion formulation of B. sacchari.

The average biomass of the non-inoculated cogongrass plants without

bahiagrass was 296.17 g/pot (Fig. 7-2). Non-inoculated cogongrass planted with

bahiagrass had an average biomass of 260.17 g/pot; thus plant competition alone did

not reduce cogongrass biomass significantly. However, significant reduction in

cogongrass biomass occurred in bahiagrass-cogongrass mixtures that were inoculated

with B. sacchari. Cogongrass subjected to plant competition and application of B.

sacchari had an average biomass of 22.83 g/pot, which was mostly the weight of the

few emergent plants and the surviving rhizomes.

March 2001-May 2001 Experiments

The application of B. sacchari to cogongrass monocultures or in a mixture

with bahiagrass significantly reduced the biomass (plant weight), rhizome production

(rhizome number), and plant height of cogongrass (Table 7-2 and Fig. 7-4).

Inoculated cogongrass (treated with B. sacchari) had an average biomass of 25.17 153

Table 7-2. Analysis of variance for cogongrass fresh weight, rhizome number and height (March- May 2001 experiments). Plant weight Rhizome number Plant Height

Source Treatment Treatment Treatment DF 3 3 3 SS 446584.7917 27864.45833 ' 12677.28023 MS 148861.5972 9288.15278 " 4225.76008 F value 35.50 31.88 45.11 Pr>F <0.0001 <0.0001 <0.0001

DF = degrees of freedom SS = sum of squares MS = mean square error 154

Fig. 7-3. Effect of application of B. sacchari on growth and biomass of cogongrass in monocultures or in a mixture with bahiagrass. Top: non-inoculated cogongrass (left) and inoculated cogongrass (right). Bottom: non-inoculated cogongrass-bahiagrass mixtures (left) and inoculated cogongrass-bahiagrass mixtures (right). Photos were taken at 2 weeks after the second inoculation. 155

Bahiagrass B. sacchari Bahiagrass + B. No treatment sacchari

Treatment

Bahiagrass B. sacchari Bahiagrass + No treatment B. sacchari Treatment

cm) 100 -,

- £ 90 D) 80 - 70 -

- C 60 CB 50 - a. m 40 - m - n 30 D> 20 - C o 10 - 0 - oo Bahiagrass B. sacchari Bahiagrass + B. No treatment sacchari

Treatment

Fig.7-4. Effect of plant competition, application of B. sacchari, and plant competition plus application of B. sacchari on the biomass (A), rhizome production (B), and plant height (C) of cogongrass. Within each graph, bars with the same letters are not significantly different at P=0.05, as determined by the Duncan's Multiple Range test. 156

400 -r

S- - Trial 1 c 350 T Trial 2 300 - a> « 250 -

(B 200 C O) 0) « 150 - M n 100 - B O) (D 50 -

a 0 - Cogongrass + B. sacchari Cogongrass Treatment

Fig. 7-5. The effect of plant competition (cogongrass) and plant competition and application of B. sacchari on the biomass of bahiagrass. Within each trial, bars with different letters are significantly different at P=0.05 as determined by the Wilcoxon Rank Sum test. 157

g/pot, produced an average of 6 rhizomes/pot, and an average plant height of 37.17 cm/pot. Untreated cogongrass had significantly greater biomass (341 .67g/pot), greater number of rhizomes (91 rhizomes/pot), and higher plant height (88.23 cm/pot). The presence of bahiagrass did not cause a reduction in the growth of cogongrass but reduced the number of cogongrass rhizomes/pot.

When cogongrass-bahiagrass mixtures were inoculated with B. sacchari, cogongrass biomass (54.83 g/pot), rhizome production (15 rhizomes/pot), and plant height (45.36 cm/pot) were all significantly lower as compared to uninoculated cogongrass planted with bahiagrass (Fig. 7- 4).

In this experiment, unlike the one that was done during the cooler months, a significant increzise in bahiagrass biomass was observed in inoculated cogongrass- bahiagrass mixtures (Fig. 7-5). The biomass of bahiagrass in the uninoculated mixtures was 94 g/pot and 192 g/pot in the two trials, while the bahiagrass biomass in inoculated mixtures was 363 g/pot and 246 g/pot, in the first and second trials, respectively.

At 2 WAI, distinct blighting was observed on inoculated cogongrass grown in monocultures or in mixtures with bahiagrass (Fig. 7-3). By 3 WAI, most of the inoculated cogongrass plants were either dead or severely blighted. Although the severely blighted plants did not show any signs of recovery, the rhizomes were able to produce green and healthy leaves between 3 and 5 WAI. Inoculated bahiagrass exhibited only minor phytotoxic damage on some of the leaf tips. 158

Discussion

The use of nonselective herbicides and soil sterilants can render the management of cogongrass difficult in situations where desirable species of grasses occur. Systems where the desirable species and the target weed are closely related provide an example of the potential value of using fungal pathogens as bioherbicides since the host specificity of such pathogens can be fully exploited (Templeton et al.,

1979).

Results from this study indicated that B. sacchari is selective on cogongrass

and that the application of B. sacchari can substantially reduce cogongrass biomass,

as well as plant height and rhizome production in cogongrass monocultures and in

cogongrass-bahiagrass mixtures. Bipolaris sacchari spores formulated in an oil

emulsion carrier caused lesions and severe blighting of cogongrass and with various

degrees of phytotoxic damage on bahiagrass plants. In experiments done in the

winter of 2000 through the spring of 2001, application of B. sacchari on bahiagrass-

cogongrass mixtures did not result in an increase or a reduction in bahiagrass growth.

The recovery of bahiagrass, a tropical grass species, from the phytotoxic damage

caused by the oil emulsion was perhaps hampered by the prevailing cooler conditions

during the course of the experiment. However, in experiments conducted from March

to May 2001, the biomass of bahiagrass was significantly increased when B. sacchari

was applied to the cogongrass-bahiagrass mixtures. A plausible explanation is that

the damage done by B. sacchari on cogongrass, which reduced its competitive ability,

combined with the favorable conditions in the greenhouse during the warmer months,

allowed bahiagrass to flourish. 159

Several studies have been done to demonstrate the mediating effect of host- specific diseases on intra- and interspecific competitive interactions (Ayres and Paul,

1990; Burdon et al., 1984; Massion and Lindow, 1986; Paul and Ayres, 1986; Paul and Ayres, 1987). Since plant pathogens can negatively affect plant growth and reproduction (Clay, 1984; Clay et al., 1989), they can be used to add additional stress

on the plant if it is grown at high density or in the presence of competition (Groves

and Williams, 1975; Alexander and Burdon, 1984; Massion and Lindow, 1986; Paul

and Ayres, 1987; Karban et al., 1989; Paul, 1989).

Burdon et al. (1984) investigated the influence of the rust Puccinia

chondrillina Bubak & Syd. on the competitive abilities of the resistant and

susceptible forms of skeletonweed {Chondrilla juncea L.). The presence of the rust

substantially reduced the dry weight of the susceptible form and increased the dry

weight of the resistant form by at least 10%. According to Groves and Williams

(1975), P. chondrillina also inhibited the ability of skeletonweed to compete in

mixture with subterranean clover (Trifolium subteraneum L.). Increased interference

from clover is considered an important aspect in the successful use of P. chondrillina

as a biocontrol agent for C. juncea in Australia (CuUen and Groves, 1977).

Paul and Ayres (1987) were able to reduce the competitive ability of common

groundsel {Senecio vulgaris L.) grown in mixture with lettuce (Lactuca sativa L.)

with the inoculation of Puccinia lagenophorae Cooke, a rust pathogen that is specific

to common groundsel. They observed that lettuce yield in plots with infected

groundsel was two to three times greater than in plots with non-infected groundsel.

In a study by Miiller-Scharer and Rieger (1998), crop losses in celeriac (Apium

graveolens var. rapaceum cv. Kojak) due to competition with 5. vulgaris was 160

significantly reduced by inoculation with P. lagenophorae. The fresh weight of

celeriac bulbs in plots with rust-infected S. vulgaris was similar to the fresh weight of

the bulbs from plots planted with celeriac alone (Muller-Scharer and Rieger, 1998).

Infection of johnsongrass {Sorghum halepense [L.] Pers.) by loose smut

{Sphacelotheca hold Jack) caused greater reduction of the rhizome length of johnsongrass and the aboveground and belowground biomass in the presence of com

{Zea mays L.) than when infected johnsongrass plants were grown alone (Massion

and Lindow, 1986).

In a study by Kadir et al. (1999), the application of Dactylaria higginsii

(Lutrell) M.B. Ellis at 10^ conidia/ml to mixed plantings of tomato (Lycopersicon

esculentus Mill.) and purple nutsedge (Cyperus rotundus L.) reduced interference

from C. rotundus and increased the yield of L. esculentum. Velvetleaf {Abutilon

theophrasti L.) suffered greater yield losses from soybean {Glycine max [L.] Merr.) in

interspecific competition in the presence of Colletotrichum coccodes (Wallr.)

Hughes. In pure velvetleaf stands, the application of C. coccodes had little impact on

seed yield of the weed. The reduction in velvetleaf seed output was attributed to the

stunting effect of the pathogen, which allowed soybean plants to grow above the

weed (Ditommaso et al., 1996). Greater deleterious effects of C. coccodes were

observed in high-density soybean-velvetleaf mixtures. This result agrees with the

observation of Massion and Lindow (1986) and Paul and Ayres (1987) that pathogens

have more detrimental effects on their host plants in crowded environments.

Potential biological control agents can be considered effective without

completely eliminating the target weed (Paul and Ayres, 1987; Watson, 1989; Watson

and Wymore, 1990). Unlike most chemical herbicides, very few biological control 161

agents can cause direct mortality of mature hosts. The inability of biological control agents to provide levels of control similar to chemical herbicides has been a deterrent in the commercialization of many biocontrol agents. However, there are situations where biocontrol agents can be used in conjunction with other existing weed control measures to control a target weed effectively. The use of plant competition and biological control agents has been suggested for weed control in range and pasture

(Bruckart and Hasan, 1991). The beneficial competitor along with the biological control agent may serve three purposes; (1) increase the rate of weed control, (2) fill the niche vacated by the weed with a plant species of value, and (3) reduce the probability that the weed will become re-established (Bruckart and Hassan, 1991).

The results presented here demonstrate the potential role of B. sacchari in an integrated approach to cogongrass management. Field trials are needed to validate the feasibility of using B. sacchari to suppress growth of cogongrass and to allow a desirable species, such as bahiagrass, to become established in natural areas. The pathogenicity of B. sacchari should also be tested on other plant species that may be used in the revegetation strategy. CHAPTER 8 SUMMARY AND CONCLUSIONS

Cogongrass, ranked as the seventh worst weed in the world, is a serious weed of several economically important crops in Asia and other tropical countries of the world

(Holm et al., 1977). In the United States, cogongrass is considered a noxious weed and is listed as one of the most invasive plant species by the Florida Exotic Pest Plant Council

(EPPC). Cogongrass thrives in natural and disturbed areas such as grasslands, roadsides, forests, recreational areas, and reclaimed mined areas in the southeastern United States

(Coile and Shilling, 1993). Cogongrass has been reported to occur in Alabama, Florida,

Georgia, Louisiana, Mississippi, South Carolina, Texas and Virginia and it continues to spread (Byrd and Bryson, 1999).

Since there is no single control measure that effectively eliminates infestations of cogongrass, integration of chemical and mechanical means has been considered as a viable approach to control cogongrass. The integrated management of cogongrass is a promising but a high-input approach to displacement of cogongrass. The cost to employ

the integrated approach precludes its use in low-maintenance and natural areas where cogongrass frequently occurs. The use of biological control agents would be ideal in

situations where high-input weed control is not feasible (Dozier et al., 1998; Shilling et

al., 1985). Biological control is also ideal in situations where a high degree of selectivity

162 163

is required to prevent nontarget injury, or where the use of chemicals is not acceptable due to environmental contamination (Auld and Morin, 1995).

A number of insects and plant pathogens have been found in association with cogongrass worldwide, some of which have been evaluated for their potential as biological control agents. However, no biological control agents are presently registered

or readily available for use in the classical or the bioherbicide approach to control

cogongrass. This situation may be due in part to the fact that none of the pathogens

tested so far was sufficiently aggressive to cause significant damage to cogongrass.

In this study, two indigenous fungal pathogens, Bipolaris sacchari (E. J. Butler)

Shoemaker and Drechslera gigantea (Heald & Wolf) Ito, were found to cause foliar

lesions and blighting on cogongrass in greenhouse tests. The disease severity caused by

each of the two pathogens ranged from 30-50%, with 50% being the maximum possible

severity. These pathogens were evaluated further for their suitability as bioherbicides for

cogongrass.

Mass-production of viable, infective, and stable propagules through inexpensive,

efficient, and reproducible methods is a major requirement for the development of a

bioherbicide (Churchill, 1982; Auld and Morin 1995). Tolerance to desiccation and

stability as a dry preparation are also required for the commercial use of bioherbicides

(Jackson et al., 1996; Jackson, 1997). In this study, production of sufficient numbers of

viable spores was achieved in the laboratory by using a solid-substrate culturing

technique, with rye grain (Secale cereale) as the substrate. Mass-produced spores on rye

grain retained their viability after drying at 30 or 40°C for 5 days and were viable for at 164

least 10 weeks after storage at low temperature (10-25°C) and low relative humidity (30-

60%).

Formulation is an important requirement for successful performance of the biological control agent in the field. Formulation is used to overcome the constraints imposed on the bioherbicide by the target weed and the environment and to enhance biocontrol efficacy and application efficiency. In greenhouse trials, the application of

spores of B. sacchari and D. gigantea formulated in an oil emulsion, composed of a

horticultural oil (Sunspray 6E) and light mineral oil, resulted in significantly higher levels

of disease even with limited dew exposure after inoculation. The even distribution of the

spores over the leaf surfaces and the phytotoxicity of the oil emulsion promoted severe

foliar blighting. Several other oils were tested as components of oil emulsions and, like

the horticultural and mineral oils, were found to have no adverse effects on the viability

of spores of B. sacchari and D. gigantea. An exception was citrus oil. Spore

formulations with canola oil, com oil, soybean oil, and sunflower oil caused disease

severities that were comparable to those caused by spore formulations applied in

Sunspray® 6E and light mineral oil. However, modifications with the vegetable oil

emulsions are needed to improve uniform dispersal of the spores in these emulsions.

Host-plant specificity has been regarded as the most important factor in the

selection of pathogens as biological control agents (Watson, 1985). It is important to

determine the host range of indigenous pathogens intended for use as bioherbicides to

avoid nontarget plant injury, such as economically important crops, and potential

conflict-of-interest issues (Weidemann, 1991). In host range studies performed by

Chandramohan (1999), only some grass species were susceptible to D. gigantea. In the case of B. sacchari, some species of economically important crop plants outside the grass family were also susceptible. The susceptibility of these non-graminaceous plants was attributed in part to the oil formulation. Application of spores with oils has been known to damage the cuticle of leaves and facilitate fungal penetration. This results in enhanced virulence of the fungal pathogens and the possible alteration of host range. Other factors that may have contributed to plant susceptibility are plant age and vigor. Nonetheless, the use of B. sacchari or D. gigantea as bioherbicides are not likely to pose the risk of

injury to nontarget plants if they are used to manage monotypic stands of cogongrass which usually occur in noncrop areas. Because the spores are formulated in oil, which improves spore adhesion on leaf surfaces, the possibility of nontarget plant injury is reduced. Nontarget plant injury is also reduced because B. sacchari and D. gigantea did not produce secondary inoculum on infected cogongrass, thus there was no inoculum to reach nontarget species.

Determination of field efficacy is one of the most important aspects of a complete evaluation of prospective weed biocontrol agents. Organisms that perform well under

controlled environments may exhibit variable activity under field conditions (Rosskopf et

al., 1999). The most frequently cited reason for variable field performance of

bioherbicides is the constraint imposed by the environment after inoculation (Greaves et

al., 1998). The duration of periods of high humidity and dew formation are the main

limiting environmental factors to the initiation and progress of disease in the field (Auld

et al., 1990; McRae and Auld, 1988). The problem with dew requirement has been

addressed by formulation of biocontrol agents with moisture-retaining materials or by the

timing of the application to coincide with periods when there is more moisture available. 166

The volume and the distribution of the applied spray are also critically important for the efficacy of a bioherbicide. In this smdy, the interaction of the spore concentration, concentration of oils in the emulsion carrier, and application volume significantly influenced the field efficacy of B. sacchari and D. gigantea. In general, a higher level of foliar blighting was produced when a higher volume of inoculum (using hand-pumped

sprayers) was applied to cogongrass. At a higher application volume (800 ml/m ), even a lower spore concentration (10^ spores/ml) and lower oil content (18% oil) contributed to the development of high levels of disease (>70%). A preliminary experiment using the

Ultra Low Volume Application (ULVA) system was performed to address the problem of

high application volume requirement. Application of B. sacchari, D. gigantea, and the

mixture of the two pathogens (with 10^ spores/ml spore concentration and 26% oil) at the

rate of 200 ml/m^, using the ULVA, resulted in >70% disease severities. Thus, it was

possible to reduce application volume with the selection of a more efficient application

system.

The use of nonselective herbicides and soil sterilants can render the management

of cogongrass difficult in situations where desirable species of grasses occur. Systems

where the desirable species and the target weed are closely related provide an example of

the potential value to use fungal pathogens as bioherbicides since the host specificity of

such pathogens is fully exploited (Templeton et al., 1979). In this study, the possibility

of B. sacchari to suppress cogongrass and to allow the establishment of a desirable

resistant species, such as bahiagrass, was investigated. Bahiagrass is a common turf

species used for soil stabilization and beautification of Florida rights-of-ways (Beard,

1980). Because cogongrass becomes established more rapidly than bahiagrass seedlings. 167

large areas planted to bahiagrass have been infested or are at risk of being infested with cogongrass (Willard and Shilling, 1990; Shilling et al., 1997). The application of spores ofB. sacchari formulated in oil to cogongrass-bahiagrass mixtures can significantly reduce cogongrass biomass, plant height, and rhizome production without causing significant damage to bahiagrass. Under favorable conditions during the warmer months,

application of B. sacchari on cogongrass-bahiagrass mixtures not only resulted in the

suppression of cogongrass but also increased the bahiagrass biomass.

The potential of Bipolaris sacchari and Drechslera gigantea as biological control

agents of cogongrass was demonstrated. These two pathogens were capable of causing

foliar damage on cogongrass. In the laboratory, mass-production of viable spores was

easily achieved through inexpensive and reproducible methods. The mass-produced

spores were amenable to drying and storage. Under field conditions, spores formulated

in oil emulsion carrier were capable of producing a severe foliar blight on cogongrass.

Like any other single control method, biological control agents may not offer the level of

control needed for long-term suppression of cogongrass. However, it is believed that

biological control can be a useful component in the integrated control approach to

management of cogongrass. REFERENCES

Abbas, H. K. and C. D. Boyette. 2000. Solid substrate formulations of the mycoherbicide Colletotrichum truncatum for hemp sesbania {Sesbania exaltata) control. Biocontrol Science and Technology 10: 291-300.

Abbas, H. K. and G. H. Egley. 1996. Influence of unrefined com oil and surface-active agents on the germination and infectivity of Altemaria helianthi. Biocontrol Science and

Technology 6 (4): 531.538.

Akobundu, I.O. 1993a. Integrated weed management techniques to reduce soil

degradation. IITA Res. 6: 1 1-16.

Akobundu, I. O. 1993b. Chemical control of cogongrass {Imperata cylindrica [L.] Raeuschel) in arable fields. Weed Sci. Soc. Am. Abst. 33:6.

Alexander, H. M. and J. J. Burdon. 1984. The effect of disease induced hy Albugo Candida (white rust) and Peronospora parasitica (downy mildew) on the survival and reproduction of Capsella bursa-pastoris (shepherd's purse). Oecologia 64: 314-318.

Alfieri, S. A., K. R. Langdon, Jr., J. W. Kimbrough, N. E. El-Gholl, and C. Wehlburg. 1994. Diseases and Disorders of Plants in Florida. Bulletin No. 14, Division of Plant

Industry, Gainesville, FL 1 1 14.p.

Amsellem, Z., A. Sharon and J. Gressel. 1991. Abolition of selectivity of two ' mycoherbicidal organisms and enhanced virulence of avirulent fungi by an invert emulsion. Phytopathology 81: 985-988

Anoka, U. A., 1. O. Akobundu, and S. N. C. Akonkwo. 1991. Effects of Gliricidia sepium (Jacq.) Steud and Leucaena leucocephala (Lam.) de Wit on growth and development of /mperato cylindrica [L.] Raeuschel. Agrofor. Syst. 16: 1-12.

Anonymous. 1978. Survey, identification and host parasite relationships of plant parzisitic nematodes associated with major economic crops in the Philippines. Technical Report NSDB Project No. 7314 Ag (2-275-53), unpublished.

Anonymous. 1995. /m/?era?a Management for Smallholders: An Extentist's Guide to Rational Imperata Management for Smallholders. Natural Resources Institute, U. K. and

Indonesian Rubber Research Institute, Jakarta, Indonesia. 56 p.

168 169

Anonymous. 2000. Bahiagrass. URL: http://www.floridaturf.com/bahia/origins.htm (11/10/00).

Aregger, E. 1992. Conidia production of the fungus Beauveria brongniartii on barley and quality evaluation during storage at 2 C. J. Invertebr. Pathol. 16: 221-226.

Aspelin, A. L. and A. H. Grube. 1999. Pesticide Industry Sales and Usage: 1996 and 1997 Market Estimates. URL World Wide Web: http://www.epa.gov.oppbead/pesticides/97pestsales/ (1 1/10/00).

Auld, B. A. 1992. Development and commercialization of biocontrol agents. Pages 269-

272 in: Proc. First Int. Weed Control Congr., J. H. Combellack, ed. AgMedia, Melbourne, Australia.

Auld, B. A. 1993. Vegetable oil suspension emulsions reduce dew dependence of a mycoherbicide. Crop Prot. 12: 477-479.

Auld, B. A. and L. Morin. 1995. Constraints in the Development of Bioherbicides. Weed Tech. 9: 638-652.

Auld, B. A., M. M. Say, H. I. Ridings, and J. Andrews. 1990. Field applications of Colletotrichum orbiculare to control Xanthium spinosum. Agric. Ecosys. Environ. 32: 315-323.

Ayres, P. G. and N. D. Paul. 1990. The effects of disease on interspecific plant competition. Aspects of Applied Biology, 24: 155-162.

Babbitt, B. 1998. Statement by Secretary of Interior on invasive alien species. Proceedings, National Weed Symposium, BLM Weed Page. April 8-10, 1998.

Bacon, P. 1986. AC 252925: a promising new compound for the control of sheet alang- alang (Imperata cylindrica [L.] Beauv.). Pages 325-339, in: Proceedings, Symposium in

Weed Science, J. V. Pancho, S. S. Sastroutomo, and S. Tjitrosemito, eds. Bogor, Indonesia: BIOTROP Special Publication No. 24.

Baker, J. M. 1970. The effects of oils on plants. Environ. Pollut. 1: 27-44.

Baker, F. W. G. 1991. Herbicide resistance. Pages 96-105 in: Tropical Grassy Weeds, F.W.G. Baker and P.J. Terry, eds. CAB International, Great Britain.

Baker, J. L. 1999. Reducing agricultural pesticide losses to surface and groundwater

resources. Pages 713-740, in: Handbook of Pest Management, J. R. Ruberson, ed. Marcel Dekker, Inc., New York. 170 ,

Baker, C. A. and J. M. S. Henis. 1990. Commercial production and formulation of microbial biocontrol agents. Pages 333-344, in: New Directions in Biological Control: Alternatives for Suppressing Agricultural Pests and Diseases. A.R. Liss, Inc., New York.

Atkins and R. H. Bannon, J. S., J. C. White, D. Long, J. A. Riley, J. Baragona, M. Cowley. 1990. Pages 305-319, in: Bioherbicide technology: an industrial perspective, R. E. Hoagland, ed. Microbes and Microbial Products as Herbicides. ACS Symp. Ser. 439. American Chemical Society, Washington, D. C.

for the Bartlett, M. C. and S. T. Jaronski. 1988. Mass production of entemogenous fungi biological control of insects. In: Fungi in Biological Control Systems, M. N. Burge, ed. Manchester University Press, Manchester, U. K.

cultivar update. Grounds Beard, J. B. 1980. St. Augustine, zoysia and bahiagrass Maintenance 15: 66-68.

Beatley, M. T. 1982. Soil erosion: its agricultural, environmental, and socioeconomic implications. Report 92, 29. Ames, lA: Council for Agric. Sci. & Tech.

Boonitee, A. and P. Ritdhit. 1984. Allelopathic effects of some weeds on mungbean plants (Vigna radiata). Proc. Trop. Weed Sci. 2:401-406.

Booth, C. and C. Prior. 1984. A new Giberella species on Imperata (Graminae). Trans. Br. Mycol. Soc. 82: 180-182.

P. J. Slininger. Bothast, R. J., D. A. Schisler, M. A. Jackson, J. E. Van Cauwenberge, and 1993. Use of pregelatinized starch and other polysaccharides for improved storage and efficacy of biocontrol agents. Pages 45-46, in: Pesticide Formulations and Application Systems: Vol. 13, P.D. Berger, B.N. Devisetty and F.R. Hall, eds. ASTM, Philadelphia, PA.

Boyette, C. D. 1982. The evaluation of Fusarium solani f.sp. cucurbita as a potential bioherbicide for controlling Texas gourd. Ph. D. Dissertation, University of Arkansas, Fayetteville.

Boyette, C. D. 1994. Unrefined com oil improved the mycoherbicidal activity of Colletotrichum truncatum for hemp sesbania {Sesbania exaltata) control. Weed Technol.

8: 526-529.

Boyette, C. D. and H. K. Abbas. 1994. Host range alteration of the bioherbicidal fungus Altemaria crassa with fruit pectin and plant filtrates. Weed Sci. 42: 487-491.

Boyette, C. D., M. A. Jackson, P. C. Quimby, Jr., W. J. Connick, Jr., N. K. Zidak, H. K. Abbas, and C. T. Bryson. 1999. Biological control of the weed hemp sesbania with Colletotrichum truncatum. In: Program Abstracts, X International Symposium on 171

Biological Control of Weeds. USDA-ARS and Montana State University, Bozeman,

MT. p. 64. Boyette, C. D., P. C. Quimby, Jr., C. T. Bryson, G. H. Egley, and F. E. Fulgham. 1993. Biological control of hemp sesbania {Sesbania exaltatd) under field conditions with Colletotrichum truncatum formulated in an invert emulsion. Weed Sci. 41: 497-500.

Boyette, C. D., P. C. Quimby, Jr., A. J. Caesar, J. L. Birdsall, W. J. Connick Jr., D. J. Daigle, M. A. Jackson, G. H. Egley, and H. K. Abbas. 1996. Adjuvants, formulations, and spraying systems for improvement of mycoherbicides. Weed Technol. 10(3): 637- 644.

Boyette, C. D., P. C. Quimby, Jr., W. J. Connick Jr., D. J. Daigle, and F. E. Fulgham. 1991. Progress in the production, formulation and application of mycoherbicides. Pages

209-222 in: Microbial Control of Weeds, D. O. TeBeest, ed. Chapman Hall, New York.

Boyette, C. D. and H. L. Walker. 1985. Factors influencing biocontrol of velvetleaf {Abutilon theophrasti) and prickly sida {Sida spinosa) with Fusarium lateritium. Weed Sci. 33:209-211.

Brian, P. W. 1976. The phenomenon of specificity in plant disease. Pages 15-22, m: Specificity in Plant Disease, R. K. S. Wood and A. Graniti, eds. Plenum Press, New York.

Bridges, D. C. 1999. General overview of weeds in crop systems. Pages 547-566, in: Handbook of Pest Management, J. R. Ruberson, ed. Marcel Dekker, Inc., New York. 842 pp.

Bright, C. 1998. Life out of Bounds: Bioinvasion in a Borderless World. W. W. Norton

& Company. New York . 287 pp.

Bruckart, W. L. and L. Hasan. 1991. Options with plant pathogens intended for classical control of range and pasture weeds. Pages 69-79, in: Microbial Control of Weeds. D.O. TeBeest, ed. Chapman and Hall, Inc., New York.

Bryson, C. T. 1985. A new food plant record for Atalopedes campestris (Boisduval)

(Hesperiidae). J. Lepid. Soc. 39: 335.

Bryson, C. T. 1987. Native butterflies accepted cogongrass ilmperata cylindrica [L.]

Beauv.) as host plant. Proc. J. Ms. Acad. Sci. 3:1.

Bryson, C. T. and R. Carter. 1993. Cogongrass, Imperata cylindrica, in the United States. Weed Technol. 7: 1005-1009.

Bunkers, G., D. Kenfield and G. A. Strobel. 1991. Production of petasol by Drechslera gigantea in liquid culture. Mycol. Res. 95(3): 347-351. :

172

Bunkers, G., D. Kenfield, G. Strobel, and F. Sugawara. 1990. Structure-activity relationships of the eremophilanes produced by Drechslera gigantea. Phytochemistry 29: 1471-1474.

Burdon, J. J., R. H. Groves, P. E. Kaye, and S. S. Speer. 1984. Competition in mixtures of susceptible and resistant genotypes of Chondrilla juncea in southeastern Australia. Journal of Applied Ecology, 18:199-203.

Burki, H., J. Lawrie, M. P. Greaves, V. M. Down, B. Juttersonke, L. Cagan, M. Vrablova, R. Ghorbani, E. A. Hassan, and D. Schroeder. 2001. Biocontrol of Amaranthus spp. in

Europe: state of the art. Biocontrol 46(2): 197-210.

Burge, M. N. 1988. The scope of fungi in biological control. Pages 3-18, in: Fungi in Biological Control Systems, M.N. Burge, ed. Manchester University Press, Manchester, U. K.

Burges, H. D. and K. A. Jones. 1998. Trends in formulation of microorganisms and future research requirements. Pages 31 1-332, in: Formulation of microbial biopesticides, H.D. Burges, ed. Kluwer Academic Publishers, The Netherlands.

Burton, G. W. 1967. A search for the origin of Pensacola bahiagrass. Econ. Bot. 21 379-382.

Busey, P. and B. J. Myers. 1979. Growth rates of turfgrass propagated vegetatively.

Agron. J. 71: 817-821.

Byrd, J. D., Jr. and C. T. Bryson. 1999. Biology, ecology, and control of cogongrass (Jmperata cylindrica [L.] Beauv.). Fact Sheet 1999-01. Mississippi Department of Agriculture and Commerce-Bureau of Plant Industry and USDA Animal and Plant Health Inspection Service-Plant Pest Quarantine.

Caster, J. 1994. Invasive alien species on D. O. T. right-of-way managed lands. In: An Assessment of Invasive Non-indigenous Species in Florida's Public Lands, D.C. Schmitz and T. C. Brown, eds. Technical Report TSS-94-00. Florida Department of Environmental Protection, Tallahassee, FL.

Gaunter, I. G. 1996. Colletotrichum caudatum, a potential bioherbicide for control of Imperata cylindrica. In: Programme and Abstracts. HI International Bioherbicide Workshop. International Bioherbicide Group, Anonymous, ed. January 19-21, 1996. Stellenbosch, South Africa.

Center, T. D., J. H. Frank, and F. A. Gray. 1997. Biological control. Pages 246-261, in: Strangers in Paradise: Impact and Management of Non-indigenous Species in Florida. D. Simberloff, D. C. Schmitz, and T. C. Brown, eds. Island Press, Washington, D. C. 173

Center, T. D., T. K. Van, M. Rayachhetry, G. R. Buckingham, F. A. Dray, S. A. Wineriter, M. F. Purcell, and P. D. Pratt. 2000. Field colonization of the Meleleuca

snout beetle (Oxyops vitiosa) in South Florida. Biological Control 19: 1 12-123.

Chambliss, C. G. 1996. Bahiagrass. Florida Cooperative Extension Service Bulletin SS- AGR-36, Gainesville, pp. 1-6.

Chandramohan, S. 1999. Multiple-pathogen strategy for bioherbicidal control of several weeds. Ph. D. Dissertation, University of Florida, Gainesville. 191 p.

Chandrasrikul, A. 1962. A preliminary host list of plant diseases in Thailand. Technical Bulletin 6. Department of Agriculture, Bangkok. 23 pp.

Chappie, A. C. and R. P. Bateman. 1997. Application systems for microbial pesticides: necessity not novelty. Pages 181-189, in: 1997 BCPC Symposium Proceedings No. 68.

Charudattan, R. 1984. Microbial control of plant pathogens and weeds. J. Georgia

Entomol Soc. 19 (Suppl. 2): 40-62.

Charudattan, R. 1985. The use of natural and genetically altered strains of pathogens for

weed control. Pages 347-372, in: Biological Control in Agricultural IPM Systems. M.A. Hoy and D.C. Herzog, eds. Academic Press Inc., Orlando, FL.

Charudattan, R. 1988. Inundative control of weeds with indigenous fungal pathogens.

Pages 88-1 10, in: Fungi in Biological Control Systems, M. N. Burge, ed. Manchester University Press, Manchester, England.

Charudattan, R. 1989a. Release of fungi: large-scale use of fungi as biological weed

control agents. In: Proc. Conf. On Risk Assessmenst in Agric. Biotechnol, J. Marois and A Browning, eds.

Charudattan, R. 1989b. Assessment of efficacy of mycoherbicide candidates. Pages 455- 464, in: "Proceedings of the VII International Symposium on Biological Control of Weeds" 1988 (E.S. Delfosse, ed.), l" Sper. Patol. Veg. (MAF), Rome, Italy. Charudattan, R. 1991. The Mycoherbicide Approach with Plant Pathogens. Pages 24-57, in: Microbial Control of Weeds, D. O. TeBeest, ed. Chapman and Hall Inc., New York.

Charudattan, R. 2001. Biological control of weeds by means of plant pathogens: Significance for integrated weed management in modem agro-ecology. Biocontrol 46: 229-260.

Charudattan, R., L. H. Walker, C. D. Boyette, W. H. Ridings, D. O. TeBeest, C. G. Van Dyke, and A. D. Worsham. 1986. Evaluation of Altemaria cassiae as mycoherbicide for sicklepod {Cassia obtusifolia) in regional field tests. Southern Coop. Ser. Bull. 317. Alabama Agric. Exp. Sta. Auburn University, Auburn, AL. 174

Chase, C. A., J. F. Gaffney, and D. G. Shilling. 1996b. Influence of glyphosate and imazapyr concentration and coverage on the efficiency of cogongrass {Imperata cylindrica [L.] Beauv.) control. WSSA Abst. 76.

Chase, C. A., D. G. Shilling, T. A. Bewick, and R. Charudattan. 1996a. Fungal isolates with potential for the biological control of cogongrass {Imperata cylindrica [L.] Beauv.). WSSA Abst. 160.

Chiang, M-Y., K. J. Leonard, and C. G. Van Dyke. 1989a. Bipolaris halepense: a new species from Sorghum halepense (johnsongrass). Mycologia 81:532-538.

Chiang, M-Y., C. G. Van Dyke, and K. J. Leonard. 1989b. Evaluation of endemic foliar fungi for potential biological control of johnsongrass {Sorghum halepense): screening and host range tests. Plant Dis. 73:459-464.

Churchill, B. W. 1982. Mass production of microorganisms for biological control. Pages 139-156 in: Biological Control of Weeds with Plant Pathogens, R. Charudattan and H. L. Walker, eds. John Wiley and Sons Inc., New York.

Ciotola, M., A. K. Watson, and S. G. Hallett. 1995. Discovery of an isolate of Fusarium oxysporum with potential to control Striga hermonthica in Africa. Weed Res. 35: 303- 309.

Clay, K. 1984. The effect of the fungus Atkinsonella hypoxolon (Clavicipitaceae) on the reproductive system and demography of the grass Danthonia spicata. New Phytologist 89: 165-175.

Clay, K., G. P. Cheplick, and S. Marks. 1989. Impact of the fungus Balansia henningsiana on Panicum agrostoides: frequency of infection, plant growth and reproduction, and resistance to pests. Oecologia 80 (3): 374-380.

Clerk, G. C. and M. F. Madelin. 1965. The longevity of conidia of three insect- parasitizing Hyphomycetes. Trans. Br. Mycol. Soc. 48: 193-209.

Coile, N. C. and D. G. Shilling. 1993. Cogongrass, {Imperata cylindrica (L.) Beauv.): A Good Grass Gone Bad! Florida Department of Agriculture and Consumer Services,

Division of Plant Industry, Botany Circular, Vol. 28. 3 p.

Colvin, D.L., J. Gaffney and D.G. Shilling. 1994. Cogongrass {Imperata cylindrica [L.] Beauv.) biology, ecology, and control in Florida. University of Florida, Institute of Food and Agricultural Services, Gainesville, FL, Circular SS-AGR-52, Weeds in the Sunshine Series. 4 p.

Connick, W. J. Jr., D. J. Daigle, and P. C. Quimby Jr. 1991. An improved invert emulsion with high water retention for mychoberbicide delivery. Weed Technol. 5- 442- 444. 175

Connick, W. J. Jr., J. A. Lewis and P. C. Quimby Jr. 1990. Formulation of biocontrol agents for use in plant pathology. Pages 345-372 in: New Directions in Biological Control: Alternatives for Suppressing Agricultural Pests and Diseases, R.R. Baker and P.E. Dunn, eds. Alan R. Liss, Inc., New York.

Cook, A. A. 1981. Diseases of Tropical and Subtropical Field, Fiber, and Oil Plants. Macmillan Publishing Co., Inc. New York, NY. 450 pp.

Cox, T. I. and D. E. Johnson. 1993. Review of research progress in Imperata cylindrica control techniques for small holder farmers. Pages 251-265, in: Proc. 3"* Tropical Weed Science Conf., S.A. Lee and K.F. Kon, eds. Malaysian Plant Protection Society, Kuala Lumpur.

Cronk, Q. and J. Fuller. 1995. Plant Invaders: The threat to natural ecosystems. Chapman & Hall, New York.

Crowder, J. P. 1974. Exotic pest plants of south Florida. South Rorida Environmental Project. Ecological Report DI-SFEP-74-23.

Cullen, J. M. and R. H. Groves. 1977. The population biology of Chondrillina juncea in southwestern Australia. Proceedings of the Ecological Society of Australia 10: 121-134.

Cunningham, J. E., C. Kuiack, and K. E. Komedant. 1990. Viability of Penicillium bilaji and Collectotrichum gloeosporioides conidia from liquid cultures. Can. J. Bot. 68: 2270- 2274.

D' Antonio, C. M. and P. M. Vitousek. 1992. Biological invasions by exotic grasses, the grass/fire cycle, and global change. Ann. Rev. Ecol. Syst. 23: 63-87.

Daigle, D. J., W. J. Connick Jr., P. C. Quimby Jr., J. P. Evans, B. Trask-Merrell and F. E. Fulgham. 1990. Invert emulsions: Carrier and water source for the mycoherbicide, Altemaria cassiae. Weed Technol. 4:327-331.

Daoust, R. A. and D. W. Roberts. 1983. Studies on the prolonged storage of Metarhizium anisopilae conidia: effect of growth substrate on conidia survival and

virulence against mosquitoes. J. Invertebr. Pathol. 41: 161-170.

Dean, C. E., D. G. Shilling, and T. R. Willard. 1988. Management of Noxious-Exotic Grasses on Highway Rights-of-Way. 99700-7352. Florida Department of Transportation.

Tallahassee, FL. 1 17 p.

DeBach, P. 1964. Biological Control of Insect Pests and Weeds. Reinhold, New York.

Dickens, R. 1974. Cogongrass in Alabama after sixty years. Weed Sci. 22:177-179. 176

Dickens, R. and G. A. Buchanan. 1971. Old weed in a new home- that's cogongrass. High. Agric. Res. 18:2.

Dickens, R. and G. A. Buchanan. 1975. Control of cogongrass with herbicides. Weed Sci. 23: 194-197.

Dickens, R. and G. M. Moore. 1974. Effects of light, temperature, KNO3, and storage on germination of cogongrass. Agronomy Journal 66: 187-188.

Dinoor, A., S. Guske, and E. Nof. 1999. Biological control of purple nutsedge {Cyperus rotundas) by pathogenic fungi. In: Abstracts, XVI International Plant Protection Congress (IPPC), July 1999. Jerusalem, Israel.

DiTommaso, A. and A. K. Watson. 1995. Impact of 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 {Ahution theophrastr)-soyhean competition in the field. Weed Science 44:924-933.

Dozier, H., J. F. Gaffney, S. K. McDonald, E. R. R. L. Johnson, and D. G. Shilling. 1998. Cogongrass in the United States: History, Ecology, Impacts and Management. Weed Technology 12: 737-743.

Drechsler, C. 1928. Zonate eyespot of grasses caused by Helminthosporium gigantea. Journal of Agricultural Research 37:473-492.

Dumas, M. T., J. E. Wood, E. G. Mitchell, and N. W. Boyonoski. 1997. Control of stump sprouting of Populus tremuloides and P. grandidentata by inoculation with Chondrostereum purpureum. Biol. Control 10: 37-41.

Egley, G. H. and C. D. Boyette. 1995. Water-corn oil emulsion enhances conidia germination and mycoherbicidal activity of Collectotrichum truncatum. Weed Sci. 43: 312-317.

Eussen, J. H. H. and M. Soerjani. 1975. Problems and control of "alang-alang" {Imperata cylindrica (L.) Beauv.) in Indonesia. Asian-Pac. Weed Sci. Soc. Fifth Conf., Bogor, Indonesia, pp. 58-65.

Falvey, J. L. 1981. Imperata cylindrica and animal production in Southeast Asia: a review. Trop. Grassl. 15:52-56.

Farr, D. F., G. F. Bills, G. P. Chamuris, and A. Y. Rossman. 1989. Fungi on Plants and Plant Products in the United States. APS Press, St. Paul. 1252 pp. 177

Fernando, W. G. D., A. K. Watson, and T. C. Paulitz. 1993. A simple technique to observe conidial germination on leaf surfaces. Mycologist 7: 188-189.

Figliola, S. S., N. D. Camper, and W. H. Ridings. 1988. Potential biological control agents for goosegrass {Eleusine indica). Weed Sci. 36:830-835.

Florida Department of Environmental Protection. 2000. Florida State Government Expenditures for Invasive Species. URL: http://www.dep.state.fl.us/stland/bapm/expenditures.htm ( 1 0/24/00).

Foudin, A. S. and O. H. Calvert. 1987. A consistent method for producing gram quantities of typical Bipolaris zeicola and B. maydis conidia. Mycologia 79(1): 1 17-122.

Frank, J. H. and E. D. McCoy. 1995. Introduction to insect behavioral ecology: the good, the bad and the beautiful: non-indigenous species in Florida. Florida Entomologist 78(1): 1-15.

Frank, J. H., E. D. McCoy, H. G. Hall, F. O'Meara, and W. R. Tschinkel. 1997. Immigration and introduction of insects. Pages 75-100, in: Simberloff D., Schmitz D. C, Brown T. C, eds. Strangers in Paradise. Island Press, Washington, D.C.

Fogal, W. H., R. K. Mittal, and G. S. Thurston. 1986. Production and quality evaluation of Beauveria bassiana for control of white spruce cone and seed insects. Information report PI-X-69, Petawa National Forestry Institute, Canadian Forestry Service, Chalk River, Ontario, Canada. ^ ^ - i I .

Gaffney, J. F. 1996. Ecophysiological and Technological Factors Influencing the Management of Cogongrass {Imperata cylindrica). Ph. D. Dissertation. University of Florida, Gainesville, FL. 128 pp.

Gaumann, E. 1950. Principles of Plant Infection. London: Crosby Lockwood. 543 p.

Greaves, M. P., P. J. Holloway, and B. A. Auld. 1998. Formulation of microbial herbicides. Pages 203-233 in: Formulation of microbial biopesticides, H. D. Burge, ed. Kluwer Academic Publishers, The Netherlands.

Greaves, M. P. and M. D. Macqueen. 1990. The use of mycoherbicides in the field. Aspects Appl. Biol. 24: 163-168. Groves, R. H. 1991. The Physiology of Grassy Weeds in Tropical Agriculture. Pages 39-51 in: Tropical Grassy Weeds, F. W. G. Baker and P. J. Terry, eds. CAB International, Great Britain.

Groves, R. H., and Williams, J. D. 1975. Growth of skeleton weed (Chondrilla juncea L.) as affected by growth of subterranean clover (Trifolium subterranean L.) and infection by Puccinia chondrillina Bubak and Syd. Australian Journal of Agricultural Research 26:975-983. 178

Guntli, D., S. Burgos, I. Kump, M. Heeb, H. A. Pfirter, and G. Defago. 1999. Biological control of hedge bindweed {Calystegia sepium) with Stagonospora convulvuli strain LA39 in combination with competition from red clover (Trifolium pratense). Biol. Control 15:252-258.

Guritno. B. S., M. Sitompul, and J. van der Heide. 1992. Reclamation of an alang-alang on an ultisol in Lampung. Agrivita 15: 87-90.

Hall, D. W. 1998. Is cogongrass really an exotic? Wildland Weeds 1:14.

Hardin, D. E. 1994. Non-indigenous species on forestry-managed lands. In: An Assessment of Invasive Non-indigenous Species in Flroida's Public Lands, D.C. Schmitz and T. C. Brown, eds. Technical Report TSS-94-00. Department of Environmental Protection, Tallahassee, PL.

Harley, K. L. S. and I. W. Porno, (eds.). 1992. Biological Control of Weeds: A Handbook for Practitioners and Students. Inkata Press, Melbourne and Sydney.

Hildebrand, D. C. and A. H. McCain. 1978. The use of various substrates for large scale production of Fusarium oxysporum f.sp. cannabis inoculum. Phytopathology 68: 1099- 1101.

Hobbs, R. J. and L. F. Huenneke. 1992. Disturbance, diversity, and invasion implications for conservation. Conserv. Biol. 6: 324-337.

Hokkanen, H. 1985. Success in classical biological control. CRC Critical Reviews in Plant Science 3(1): 35-72.

Holcomb, G. E. 1982. Constraints on disease development. Pages 61-71 m: Biological Control of Weeds With Plant Pathogens, R. Charudattan and H. L. Walker, eds. John Wiley &. Sons, New York.

Holliday, P. 1980. Fungus Diseases of Tropical Crops. Cambridge University Press, Great Britain. 607 pp.

Holm, L. G., D. L. Plucknett, J. V. Pancho, and J. P. Herberger. 1977. The World's Worst Weeds: Distribution and Biology. University Press of Hawaii, Honolulu. 609 pp.

Hooker, A. L., D. R. Smith, S. M. Lim, and J. B. Beckett. 1970. Reaction of seedlings with male-sterile cytoplasm to Helminthosporium maydis. Plant Dis. Rep. 54: 708-712.

Hubbard, C. E., D. Brown, A. P. Gray, and R. O. Whyte. 1944. Jmperata cylindrica: Taxonomy, Distribution, Economic Significance and Control. Imperial Agricultural Bureau Joint Publication 7:1-63. 179

Hussain, F. N. Abidi, S. Ayaz and A-U-R Saljoqi. 1992. Allelopathic suppression of wheat and maize seedling growth by Imperata cylindrica (L.) Beauv. Sarhad Journal of Agriculture 8(4): 433-439.

Imaizumi, S., T. Nishino, K. Miyabe, T. Fujimori, and M. Yamada. 1997. Biological control of annual bluegrass {Poa annua L.) with a Japanese isolate of Xanthomonas campestris pv. poae (JT-P482). Biol. Control 8:7-14.

Inderjit and K. M. M. Dakshini. 1991. Investigations on some aspects of chemical ecology of cogongrass, Imperata cylindrica (L.) Beauv. J. Chem. Ecol. 17: 343-352.

IRRI (International Rice Research Institute). 1988. International Rice Research Newsletter 13(4): 30-33, 37. 7314 Ag (2-275-53), Unpublished.

Jackson, A. 1997. Optimizing nutritional conditions for the liquid culture production of effective fungal biological control agents. Journal of Industrial Microbiology and Biotechnology 19: 180-187.

Jackson, M. A. 1997. Optimizing nutritional conditions for the liquid culture production of effective fungal biological control agents. J. Ind. Microbio. Biotechnol. 19:180-187.

Jackson, M. A. and R. J. Bothast 1990. Carbon concentration and carbon-to-nitrogen ratio influence submerged culture conidiation by the potential bioherbicide Colletotrichum truncatum NRRL 13737. Appl. Environ. Microbiol. 56: 3435-3438.

Jackson, M. A. and D. A. Schisler. 1992. The composition and attributes of Colletotrichum truncatum spores altered by the nutritional environment. Appl. Environ. Microbiol. 58: 2260-2265.

Jackson, M. A., D. A. Schisler, P.J. Sliniger, C. D. Boyette, R.W. Silman, and R. J. Bothast. 1996. Fermentation strategies for improving the fitness of a bioherbicide. Weed Technol. (10) 645-650.

James, C. 1971. A Manual of Assessment Keys for Plant Diseases. Canada Dept. of

Agriculture, Ottawa, Canada; American Phytopthological Society, St. Paul, MN. 78 p.

Johnson, R. 1976. Genetics of host-parasite interactions. Pages 45-62, in: Specificity In Plant Disease, R. K. S. Wood and A. Graniti, eds Plenum Press, New York.

Jones, K. A. 1998. Spray application criteria. Pages 367-375 in: Formulation of microbial biopesticides, H.D. Burges, ed. Kluwer Academic Publishers, The Netherlands.

Kadir, J. B. and R. Charudattan. 2000. Dactylaria higginsii, a fungal bioherbicide agent fox Tpuvp[& nvXsodgQ {Cyperus rotundus). Biol. Control 17: 113-124. 180

Kadir, J. B., R. Charudattan, W. M. Stall, and T. A. Bewick. 1999. Effect of Dactylaria higginsii on interference of Cyperus rotundus with L. esculentum. Weed Sci. 47: 682- 686.

Karban, R., A. K. Brody, and W. C. Schnathorst. 1989. Crowding and a plant's ability to defend itself against herbivores and diseases. American Naturalist 134: 749-760.

Kenfield, D., F. Sugawara, F., G. Bunkers, Y. H. Wu, G. Strobel, Y. Fu, and J. Clardy. 1989. Gigantenone, a novel sesquiterpene phytohormone mimic. Experientia 45: 900- 902.

Klein, T.A. and B. A. Auld. 1995. Influence of spore dose and water volume on a mycoherbicide's efficacy in field trials. Biological Control 5:173-178.

Klose, N. 1950. America's Crop Heritage. The Iowa State College Press, Ames, Iowa. 156 pp.

Kroschel, J., D. Muller-Stover, A. Elzein and J. Sauerbom. 1999. The development of mycoherbicides for the management of parasitic weeds of the genus Striga and Orobanche- a review and recent results. In: Program and Abstracts, X International Symposium on Biological Control of Weeds. USDA-ARS and Montana State

University, Bozeman, MT. p. 24.

Kurdila, J. 1995. The introduction of exotic species into the United States: there goes the neighborhood. Environmental Affairs 16: 95-118.

Langeland, K. A. and K. C. Burks. 1998. Identification and biology of non-native plants in Florida. University of Florida, Gainesville, FL. 165 p.

LeBaron, H. M. and J. Gressel (eds.). 1982. Herbicide Resistance in Plants. John Wiley & Sons, Inc., New York.

LeBaron, H. M. and J. McFarland. 1990. "Overview and prognosis of herbicide resistance in weeds and crops", Proceedings of the American Chemical Society Symposium on Fundamental and Practical Approaches to Combating Resistance, ACS Symposium Series, ACS Books, Washington, D. C.

Lee, S. A. 1985. Bud development in the rhizomes of Imperata cylindrica [L.] Beauv. after glyphosate treatment. MARDI Res. J. 14: 39-45.

Legget, M. E. and S. C. Gleddie. 1995. Developing biofertilizer and biocontrol agents that meet farmer's expectations. Adv. Plant Pathol. 1 1:59-74.

Leth, V. and C. Andreasen. 1999. Phomopsis cirsii: A promising control agent for Cirsium arvense. In: Program Abstracts, X International Symposium on Biological

Control of Weeds. USDA-ARS and Montana State University, Bozeman, MT. p. 1 16. 181

Lippincott, C. L. 1997. Ecological consequences of Imperata cylindrica (Cogongrass) invasion in Florida sandhill. Ph. D. Dissertation, University of Florida, Gainesville. 164

P-

Livingston, R. S. and R. P. Scheffer. 1984. Selective toxins and analogs produced by Helmithosporium sacchari. Plant Physiol. 76: 96-102.

Macko, v., T. J. Wolpert, W. Acklin, and D. Arigoni. 1989. Biological activities and structural variants of host-selective toxins from Cochliobolus victoriae. Pages 31-41, in: Phytotoxins and Plant Pathogenesis, A. Graniti, R. D. Durbin, and A. Ballio, eds. Springer-Verlag, Berlin.

MacLagan Gorrie, A. 1950. Dry zone afforestration and agriculture development in Ceylon. British Agricultural Bulletin 3:70.

Malecki, R. A., B. Blossey, S. D. Hight, D. Schroeder, L. T. Kok, and J. R. Coulson. 1993. Biological control of purple loosestrife. Bioscience 43: 680-686.

Makowski, R. M. D. 1993. Effect of inoculum concentration, temperature, dew period and plant growth stage on disease of round-leaved mallow and velvetleaf by

Colletotrichum gloeosporiodes f. sp. malvae. Phytopathology 83: 1229-1234.

Mallory-Smith, C. A., D. C. Thill, and M. J. Dial, 1990. Identification of sulfonylurea herbicide resistant prickly lettuce {Lactuca serriola). Weed Tech. 4: 163.

Manyong, V. M., A. V. Houndekon, A. Govan, M. V. Versteeg, and F. van der Pol. 1996. Pages 86-89 in: Determinants of Adoption for a Resource Management Technology: The Case of Mucuna in Benin Republic. Proc. International Conference of Advances in Agricultural and Biological Environment Engineering Conf., Beijing 15-19 August. China Agricultural University Press, Beijing.

Massion, C.L. and S. E. Lindow. 1986. Effects of Sphacelotheca hold infection on morphology and competitiveness of Johnsongrass {Sorghum halepense). Weed Science 34: 883-888.

Masuzawa, T., H. Suwa, and F. Nakasuji. 1983. Differences of oviposition preference and survival rate of two skipper butterflies Pamara guttata and Pelopidas mathias (Lepidoptera: Hesperiidae) on rice plant and cogongrass. New Entomol. 32: 1-10.

McCoy, E. E. and C. W. Carver. 1941. A method for obtaining spores of the fungus

Beauveria bassiana (Bals) Vuill. in quantity. J. N.Y. Entomol. Soc. 45:205-210.

McDonald, S. K., D. G. Shilling, T. A. Bewick, D. Gordon, D. Hall, and R. Smith. 1996. Factors influencing cogongrass, Imperata cylindrica (L.) Beauv., dispersion, establishment, and persistence. Weed Sci. Soc. Am. Absts. 36:46. 182

McEvoy, P. B. and N. T. Rudd. 1993. Effects of vegetation disturbances on insect biological control of tansy ragwort, Senecio jacobea. Ecol. Appl. 3: 682-698.

McRae, C. F. and B. A. Auld. 1988. The influence of environmental factors on anthracnose of Xanthium spinosum. Phytopathology 78:11 82- 1 1 86.

McWhorter, C. G. and W. L. Barrentine. 1988. Spread of parrafinic oil on leaf surfaces of johnsongrass {Sorghum halepense). Weed Sci. 36: 1 11-1 17.

Meredith, D.S. 1963. Ann. Appl. Biol. 51:29 (on banana, general; 42:476).

Misra, A. P. 1973. Helminthosporium species occurring on cereals and other graminae. Catholic Press, Dholi Muzaffarpur (Bihar), India.

Moore, D., O. K. Douro-Kpindou, N.E. Jenkins and C.J. Lomer. 1996. Effects of moisture content and temperature on storage of Metarhizium flavoviridae conidia. Biocontrol Science and Technology 6(1): 51-61.

Morin, N. 1995. Vascular plants of the United States. Pages 200-205, in: Our Living Resources: A Report to the Nation on the Distribution, Abundance, and Health of U.S. Plants, Animals and Ecosysytems, LaRoe, E. T., Farris, G. S., Puckett, C. E., Doran, P.

D. Mac, M. J., eds. U. S. Department of the Interior, National Biological Service, , Washington, D. C.

Morin, L. A., A. K. Watson, and R. D. Reeleder. 1990a. Effect of dew, inoculum density and spray additives on infection of field bindweed by Phomopsis convolvulus. Can. J. Plant Pathol. 12: 48-56.

Morin, L., A. K. Watson, and R. D. Reeleder. 1990b. Production of conidia by

Phomopsis convolvulus. Can. J. Microbiol. 36: 86-91.

Morris, M. J., A. R. Wood, and A. Den Breeyen. 1998. Development and registration of

a fungal inoculant to prevent regrowth of cut wattle tree stump in South Africa , and a brief overview of other bioherbicide projects currently in progress. 'TV International Bioherbicide Workshop Programme and Abstract, 6-7 August 1998". Univ. of Strathclyde, Glasgow, Scotland. P. 15

Morse, L. E., J. T. Kartesz, L. S. Kutner. 1995. Native vascular plants. Pages 205-209,

in: Our Living Resources: a Report to the Nation on the Distribution, Abundance, and Health of U. S. Plants, Animals and Ecosystems, LaRoe, E. T., Farris, G. S., Puckett, C.

E. , Doran, P. D., Mac, M. J., eds. U. S. Department of the Interior, National Biological Service, Washington, D. C. 183

Mortensen, K. 1988. The potential of an endemic fungus, Co//erofric/ium gloeosporioides, for biological control of roundleaved mallow (Malva pusilla) and velvetleaf {Abutilon theophrasti). Weed Sci. 36: 473-478.

Mortensen, K. and Makowski, R.M.D. 1989. Field efficacy at different doses of Colletotrichum gloeosporiodes f.sp. malvae as a bioherbicide for round-leaved mallow

(Malva pusilla). Pages 523-530, in: "Proceedings of the Vn International Symposium on Biological Control of Weeds" 1988 (E.S. Delfosse, Ed.), l" Sper. Patol. Veg. (MAP), Rome, Italy.

Miiller-Scharer, H. and S. Rieger. 1998. Epidemic spread of the rust fungus Puccinia

lagenophorae and its impacts on the competitive ability of Senecio vulgaris in celeriac during early development. Biocontrol Sci. and Technol. 8: 59-72.

Munyaradzi, S. T., M. Campbell, and M. N. Surge. 1990. The potential for bracken control with mycoherbicide formulations. Aspects Appl. Biol. 24: 169-175.

Myers, R. L. 1990. Problems, prospects, and strategies for conservation. In:

Ecosystems in Florida, R. L. Myers and J. J. Ewel, eds. University of Central Florida Press, Orlando.

Office of Technology Assesment. 1993. Harmful Non-indigenous Species in the United States. U. S. Congress/OTA-F-565. U. S. Congress, Office of Technology Assessment, Washington, D. C.

Oku, H., T. Shiraiski, and S. Ouchi. 1979. The role of phytotalexins in host-parasite specificity. Pages 317-333, in: Recognition and Specificity in Plant Host-Parasite

Interactions, J. M. Daly and I. Uritani, eds. Japan Sci. Soc. Press, Tokyo.

Ormeno-Nunez, J. R. D. Reeleder, and A. K. Watson. 1988. A foliar disease of field bindweed (Convolvulus arvensis) caused by Phomopsis convolvulus. Plant Dis, 72: 338- 342.

Patterson, D. T., E. P. Flint, and R. Dickens. 1980. Effects of temperature, photoperiod, and population source on the growth of cogongrass (Imperata cylindrica). Weed Sci. 28:505-509.

Patterson, D. T., E. E. Terrell, and R. Dickens. 1979. Cogongrass in Mississippi. Miss. Agric. For. Exp. Stn. Res. Rep. 46(6): 1-3.

Paul, N. D. 1989. The effects of Puccinia lagenophorae on Senecio vulgaris in competition with Euphorbia peplus. Journal of Ecology 77: 552-564.

Paul, N. D., and Ayres, P. G. 1987. Effects of rust infection of Senecio vulgaris in competition with lettuce. Weed Research 27: 431-441. 184

Paul, N. D., and Ayres, P. G. 1986. Interference between healthy and rust-infected groundsel {Senecio vulgaris L.) within mixed populations of different densities and proportions. New Phytol. 104: 257-269.

Pendleton, R. L. 1948. Cogongrass, Imperata cylindrica in the western hemisphere. J. of Agr. 40: 1047-1049.

Persley, G. J. 1973. Naturally occurring alternative hosts of Xanthomonas albineans in Queensland. Plant Dis. Reprt. 57: 1040-1042.

Pfirter, H. A., H. Ammon, D. Guntli, M. P. Greaves, and G. Defago. 1997. Towards the management of field bindweed {Convolvulus arvensis) and hedge bindweed {Calystegia sepium) with fungal pathogens and cover crops. Int. Pest Manage. Rev. 2:61-69.

Pimm, S. L. and M. E. Gilpin. 1989. Theoretical issues in conservation. Pages 287-305, in: Perspectives in Ecological Theory, J. Roughgarden, R. M. and S. A. Levin eds. Princeton University Press, New Jersey.

Pimentel, D. L. 1991. Handbook on Pest Management in Agriculture. Volumes 1,2, and 3. CRC Press, Boca Raton, PL.

Pimentel, D. 1993. Habitat factors in new pest invasions. Pages 165-181, m: Evolution of Insect Pests-Pattems of Variation. Kim, K.C. and McPheron, B.A., eds. John Wiley & Sons, New York. Pimentel, D., L. Lach, R. Zuniga, and D. Morrison. 1999. Environmental and economic costs associated with non-indigenous species in the United States. URL: http://www.news.comell.edu/releases/Jan99/species_costs.htm.

Pimentel, D., M. S. Hunter, J. A. Lagro, R. A. Efronymson, J. C. Landers, F. T. Mervis, C. A. McCarthy, and A. E. Boyd. 1989. Benefits and risks of genetic engineering in agriculture. Bioscience 39: 606-614.

Populer, C. 1978. Changes in host specificity with time. Pages 239-262, in: Plant

Disease: An Advanced Treatise, Vol. H, J. G. Horsfall and E. B. Cowling, eds. Academic Press, New York.

Potyka, I. 1996. Emulsion-formulation of microbial herbicides, Ph. D.Thesis, University of Bristol.

Powles, S.B. and Howat, P.D. 1990. Herbicide resistant weeds in Australia, Weed Technology 4: 178-185.

Prasad, R. 1994. Influence of several pesticides and adjuvants on Chondrostereum purpureum- a bioherbicide agent for control of forest weeds. Weed Technol. 8: 445-449. 185

Primiani, M. M., J. C. Cotterman and L. L. Saari. 1990. Resistance of kochia (Kochia scoparia) to sulfonylurea and imidazolinone herbicides. Weed Tech. 4:169.

Quarin, C. L., B. L. Burson, and G. W. Burton. 1984. Cytology of intra- and interspecific hybrids between two cytotypes of Paspalum notatum and P. cromyorrhizon. Bot. Gaz. 145: 420-426.

Quimby, P. C. Jr., F. E. Fulgham, C. D. Boyette, and W. J. Connick Jr. 1988. An invert emulsion replaces dew in biocontrol of sicklepod- a preliminary study. Pages 264-270, in: Pesticide Formulations and Application Systems, A. Hovde and G. B. Beestman, eds. ASTM-STP 980D, Vol. 8. American Society for Testing and Materials. Philadelphia.

Radosevich, S., J. Holt, and C. Ghersa. 1997. Weed Ecology: Implications for 2"** Management, ed. John Wiley & Sons, Inc., New York, N. Y. 589 pp.

Randall, J. M. 1996. Weed control for the preservation of biological diversity. Weed Technology 10: 370-383.

Randall, J. M. and J. Marinelli, editors. 1996. Invasive plants: Weeds of the global garden. Brooklyn Botanical Garden, Brooklyn, New York.

Ravenell, D. I., and C. G. Van Dyke. 1986. Efficacy and host range of Bipolaris setariae as a potential biocontrol pathogen of broadleaf signal grass (Brachiaria platyphylla [Griseb.] Nash.). WSSA Abst. 26:54.

Rhodes, D. J. 1993. Formulation of biological control agents. Pages 41 1-439, in: Exploitation of Microorganisms, D. G. Jones, ed. Chapman Hall, London.

Rhodes, D. J., K. A. Powell, M. P. Macqueen, and M. P. Greaves. 1990. Controlled delivery of biological control agents. In: Controlled Delivery of Biological Control Agents, R.M. Wilkins, ed. Taylor and Francis.

Rice, E. L. 1995. Biological Control of Weeds and Plant Diseases: Advances in Applied Allelopathy. University of Oklahoma Press, Norman, Publishing Division of the University. USA.

Ridings, W. H. 1986. Biological control of stranglervine in citrus- a reseacher's view. Weed Sci. 34 (Suppl.l): 31-32.

Ross, M. and C. Lembi. 1983. Applied weed science. Minneapolis, Minnesota: Burgess Publishing Company. Ph. D. Thesis Univ. of Florida, Gainesville, FL, USA. 187 pp.

Rosskopf, E. N. 1998. Evaluation of Phomopsis amaranthicola sp. nov. as a biological control agent for Amaranthus spp. Ph. D. Dissertation. University of Florida, Gainesville, FL. 190 pp. 186

Rosskppf, E. N., R. Charudattan and J. B. Kadir. 1999. Use of Plant Pathogens in Weed

Control. Pages 891-917, in: Handbook of Biological Control. San Diego, CA.: Academic Press.

Ryan, G. F. 1970. Resistance of common groundsel to simazine and atrazine. Weed Science 18: 614.

Ryerson, K. A. 1967. The history of plant exploration and introduction into the United States Department of Agriculture. In: Proceedings of the International Symposium on Plant Production. Escuela Agricola Panamericana, Tegucigalpa, Honduras.

Sajise, P. E. 1980. Alang-alang {Imperata cylindrica (L.) Beauv.) and upland agriculture. BIOTROP 5: 35-46.

Salmon, J. T. 1954. A new polyvinyl alcohol mounting medium. The Microscope 10: 66-68.

Sankaran, K. V. and B. C. Sutton. 1991. Myrellina imperatae sp. nov. on Imperata from Papua New Guinea and Australia. Mycol. Res. 95: 1021-1022. SAS Institute. 1988. SAS/STAT User's Guide, release 6.12, SAS Institute, Gary, N.C.

Scheffer, R. P. 1976. Host-specific toxins in relation to pathogenesis and disease resistance. Pages 247-269, in: Physiological Plant Pathology, R. Heitefuss and P. H. Williams, eds. Springer-Verlag, Berlin.

Scheffer, R. P. 1989. Ecological consequences of toxin production by Cochliobolus and related fungi. Pages 285-300, in: Phytotoxins and Plant Pathogenesis, A. Graniti, R. D. Durbin, and A. Ballio, eds. Springer-Verlag, Berlin.

Scheffer, R.P. 1991. Role of toxins in evolution and ecology of plant pathogenic fungi. Experientia 47: 804-811.

Scheffer, R.P. and R.S. Livingston. 1980. Sensitivity of sugarcane clones to toxin from Helminthosporium sacchari as determined by electrolyte leakage. Phytopathology 70: 400-404.

Schisler, D. A., K. M. Howard, and R. J. Bothast. 1991. Enhancement of disease caused by Colletotrichum truncatum in Sesbania exaltata by co-inoculating with epiphytic bacteria. Biol. Control 1: 261-268.

Schroeder, D. 1983. Biological Control of Weeds. Pages 41-78, in: Recent Advances in Weed Research, W. W. Fletcher, ed. Commonw. Agr. Bur., Famham Royal, England.

SEAWIC (Southeast Asian Weed Information Center). 1988. Annotated bibliography. SEAMEO-BIOTROP Center, Bogor, Indonesia. 2: 1-37. 187

Shabana, Y. M., M. A. Elwakil, and R. Charudattan. 1999. Development of Altemaria eichomiae Nag Raj & Ponnappa for biological control of water hyacinth in Egypt. In: Proceedings of the First Regional Symposium for Applied Biological Control in Mediterranean Countries, M. Canard and V. B. Amaouty, eds. Cairo, Egypt, pp. 211- 215.

Shamoun, S. F., and W. E. Hintz. 1998. Development and registration of Chondrostereum purpureum as a mycoherbicide for hardwood weeds and conifer reforestration sites and utilities rights-of-way. "IV International Bioherbicide Workshop Programme and Abstracts, 6-7 Aug., 1998." P. 14. Univ. of Strathclyde, Glasgow, Scotland.

*

Shilling, D. G., T. A. Bewick, J. F. Gaffney, S. K. McDonald, C. A. Chase, and E. R. R. L. Johnson. 1997. Ecology, Physiology, and Management of Cogongrass {Imperata cylindrica). Final Report. Rorida Institute of Phosphate Research. 128 p.

Shilling, D. G., J. F. Gaffney, and P. Waldrop. 1995. Cogongrass: problem and solutions. Alabama's Treasured Forests 3: 8-9.

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.

Silman, R. W., T. C. Nelsen, and R. J. Bothast. 1991. Comparison of culture methods for production of Colletotrichum truncatum spores for use as a mycoherbicide. FEMS Microbiology Letters 79: 69-74.

Simberloff, D. 1996. Impacts of Introduced Species in the United States. Consequences. URL: World Wide Web: http://www.gopher.gcrio.org/CONSEOUENCES/vol2no2/article2.html (9/6/00).

Simberloff, D., D. C. Schmitz, and T. C. Brown (eds.). 1997. Strangers in Paradise: Impact and Management of Non-indigenous Species in Florida. Island Press, Washington, D. C.

Small, J. K. 1933. Manual of the Southeastern Flora. University of North Carolina

Press, Chapel Hill, N. C. p. 39

Smith, R. J., Jr. 1982. Integration of microbial herbicides with existing pest management programs. Pages 189-203, in: Biological Control of Weeds with Plant Pathogens, R. Charudattan and H. L. Walker, eds. Wiley, New York.

Smith, R. J., Jr. 1986. Biological control of northern jointvetch in rice and soybeans- a researcher's view. Weed Sci. 34 (Suppl. 1): 17-23.

Soerjani, M. 1970. Alang-alang, Imperata cylindrica (L.) Beauv. (1812). Pattern of growth as related to its problem of control. BIOTROP 1. p. 88. 188

Sonoda, R. M., B. M. Turner, and T. S. Schubert. 1992. Bipolaris sacchari on Panicum ma;ci>nMm in Horida. Plant Disease 77: 101.

Sriyani, N. 1992. Glyphosate absorption, translocation, and rhizome bud kill in cogongrass {Imperata cylindrica). Proc North Cent. Weed Sci. Soc. 47: 108.

Steiner, G.W. and G.A. Strobel. 1971. Helminthosporoside, a host specific toxin from

Helmithosporium sacchari. J. Biol. Chem. 246: 4350-4357.

Steinhaus, E. A. 1960. The duration of viability and infectivity of certain insect pathogens. J. Invertebr. Pathol. 2:225-229.

Stowell, L. J. 1991. Submerged fermentation of biological herbicides. Pages 225-261 in: Microbial Control of Weeds, D. O. TeBeest, ed. Chapman and Hall Inc., New York. Tabor, P. 1952. Comments on cogon and torpedo grasses: A Challenge to Weed

Workers. Weeds 1: 374-375.

Tabor, P. 1949. Cogongrass, Imperata cylindrica (L.) Beauv., in the southeastern United

States. Agron. J. 41:270.

Tanner, G. W., J. M. Wood, and S. A. Jones. 1992. Cogongrass (Imperata cylindrica) control with glyphosate. Fla. Sci. 55: 1 12-115.

TeBeest, D. O. 1985. Techniques for testing and evaluating plant pathogens for weed control. J. Agric. Entomol. 2: 98-134.

TeBeest, D. O. 1988. Techniques for testing and evaluating plant pathogens for weed control. J. Agric. Entomol. 2: 98-134.

TeBeest, D. O., and G. E. Templeton. 1985. Mycoherbicides: progress in the biological control of weeds. Plant Dis. 69: 6-10.

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: 637-57.

Tempany, H. 1951. Imperata grass, a major menace in wet tropics. World Crops 3: 143- 146.

Templeton, G. E. 1982. Biological Herbicides: Discovery, Development, Deployment. Weed Sci. 30: 430-433.

Templeton, G.E., D. O. TeBeest, and Smith, R. J., Jr. 1979. Biological weed control with mycoherbicides. Annu. Rev. Phytopathol. 17:301-310. 189

Templeton, G.E., D. O. TeBeest, and Smith, R. J., Jr. 1984. Biological weed control in rice with a strain of Colletotrichum gloeosporiodes (Penz.) Sacc. used as a mycoherbicide. Crop Protection 3: 409-422.

Thompson, D. Q., R. L. Stuckey, and E. B. Thompson. 1987. Spread, impact and control of purple loosestrife {Lythrum salicaria) in North American wetlands. U. S. Fish and

Wildlife Serv. Res. Rep. 2.

Townson, J. K. and R. Butler. 1990. Uptake, translocation and phytotoxicity of imazapyr and glyphosate in Imperata cylindrica [L.] Raeuschel: effect of herbicide concentration, position of deposit and two methods of direct contact application. Weed Res. 30: 235-343.

Townson, J. K. and C. E. Price. 1987. Tropical weed control: an approach to optimizing herbicide performance with very low application volume. Aspects Appl. Biol. 14: 305- 306. United States Bureau of the Census. 1998. Statistical Abstract of the United States. 1996. 200*^ ed. US Bureau of Census, U. S. Government Printing Office, Washington, D. C.

Van Tuat N., H. A. Minh Trung, S. D. Hetherington, and B. A. Auld. 1998. Potential bioherbicide for Echinochloa in Vietnam. In: Programme and Abstracts, FV International Bioherbicide Workshop, August 1999, Glasgow. University of Strathclyde,

Glasgow, Scotland, p. 45.

Vaysierre, P. 1957. Les mauvaises herbes en Indo-Malasie. Journal d' agriculture tropical et de botanique applique 4(9-10): 392-401.

Versteeg, M. V. and V. Koudokpon. 1990. Mucuna helps control Imperata in southern Benin. West Afr. Farming Syst. Res. Network Bull. 7: 7-8.

Vincent, A. C. and R. Charudattan. 1999. Effects of formulations of Myrothecium roridum Tode ex. Fr. and Cercospora rodmanii Conway on waterhyacinth {Eichomina crassipes [Mart.] Solms-Laub.) under greenhouse and field conditions. WSSA Abstracts 39:71.

Vitousek, P. M., C. M. D'Antonio, L. L. Loope, and R. Westbrooks. 1996. Biological invasions as global environmental change. American Scientist 84: 468-478.

Vitousek, P. M., C. M. D'Antonio, L. L. Loope, M. Rejmanek, and R. Westbrooks. 1997. Introduced species: a significant component of human-caused global change. New Zeziland Journal of Ecology 21(1): 1-16.

Vurro, M., A. Evidente, M. C. Zonno, R. Pengue, P. Montemurro, and B. Auld. 1999. Phytotoxic metabolites produced by Drechslera avenacea, a potential mycoherbicide for Avena fatua In: Program Abstracts, X International Symposium on Biological Control of Weeds. USDA-ARS and Montana State University, Bozeman, MT. p. 71. 190

Waage, J. K. and D. J. Greathead. 1988. Biological Control: Challenges and Opportunities. Philosophical Transactions of the Royal Society of Lx)ndon. B. 318: 111- 128.

Wagner, W. H. 1993. Problems with biotic invasions: a biologist's viewpoint. Pages 1-8 in: Biological Pollution: The Control and Impact of Invasive Exotic Species. B. N. McKnight, ed. Indiana Academy of Sciences, Indianapolis, Indiana.

Walker, H. L. 1982. Seedling blight of sicklepod caused by Altemaria cassiae. Plant Dis. 66: 426-428.

Walker, H. L. 1980. Altemaria macrospora as a potential biocontrol agent for spurred anoda: production of spores for field studies. U.S. Department of Agriculture, Science and Education Administration Advances in Agricultural Technology.

Walker, H. L. and J. A. Riley. 1982. Evaluation of Altemaria cassiae for the biocontrol of sicklepod {Cassia obtusifolia). Weed Sci. 30: 651-654.

Wapshere, A. J. 1974. A strategy for evaluating the safety of organisms for biological

weed control. Ann. Appl. Biol. 77: 201-21 1.

Wapshere, A. J. 1982. Biological control of weeds. Pages 47-56, in: Biology and Ecology of Weeds, W. Holzner and N. Numata, eds. Dr. W. Publ., The Hague.

Wapshere, A. J. 1983. Problems in the use of plant biochemistry for establishing the safety of biological control agents for weeds: the Chondrilla and EchiumlHeliotropium cases. Entomophaga 28: 287-294.

Watson, A. K. 1985. Host specificity of plant pathogens in biological weed control.

Pages 577-586, in: VI Int. Symp. Biol. Control Weeds, E. S. Delfosse, ed. Agric. Can., Can. Gov. Publ. Ctr., Ottawa.

Watson, A. K. 1989. Current advances in bioherbicide research. Brighton Crop Protection Conference- Weeds. 8B-3: 987-996.

Watson, A.K. and Wymore, L.A. 1990. Identifying limiting factors in the biocontrol of weeds. Pages 305-316, in: New Directions in Biological Control, R. Baker, and P. Dunn, eds. UCLA Symposium of Molecular and Cellular Biology, New Series 112. Alan R. Liss, New York.

Watson, L. and M. J. Dallwitz. 1992. Grass genera of the world: descriptions. Illustrations, identification, and information retrieval; including synonyms, morphology, anatomy, physiology, phytochemistry, cytology, classification, pathogens, world and

local distribution, and references. URL, http:// www.keil.ukans.edu/delta/ . 191

Watson, V. H. and B. L. Burson. 1985. Bahiagrass, carpetgrass, and dallisgrass in Forages: the science of grassland agriculture. Iowa State University Press, Ames. Pp. 255-257.

Weber, F. J., J. Tramper and A. Rinzema. 1999. A simplified material and energy balance approach for process development and scale-up of Coniothyrium minitans conidia production by solid-state cultivation in a packed-bed reactor. Biotechnology and Bioengineering 65(4): 447-458.

Weidemann, G. J. 1991. Host-range testing: safety and science. Pages 83-96, m: Microbial Control of Weeds, D. O. TeBeest, ed. Chapman and Hall, New York.

Westbrooks, R. 1998. Invasive plants, changing the landscape of America: Fact book. Washington, D. C: Federal Interagency Committee for the Management of Noxious and Exotic Weeds (HCMNEW). 109 pp.

Westbrooks, R.G. and R.E. Eplee. 1989. Federal Noxious Weeds in Florida. Proc. South. Weed Sci. Soc. 42: 316-321.

Whipps, J. M. and M. Gerlagh. 1992. Biology of Coniothyrium minitans and its potential for use in disease biocontrol. Mycol. Res. 96:897-907.

Whisenant, S. G. 1990. Changing fire frequencies on Idaho's Snake River Plain: Ecological and Management Implications. The Station. Nov. 1990 (276). Ogden, Utah: General Technical Report INT-U. S. Department of Agriculture, Forest Service, Intermountain Research Station.

Wicks, G. A., O. C. Bumside, and W. L. Felton. 1995. Mechanical weed management.

Pages 51-99 in: Handbook of Weed Management, A. E. Smith, ed. Marcel Dekker, Inc., New York.

Wilcut, J. W., R. D. Dute, B. Truelove, and D. E. Davis. 1988a. Factors limiting the distribution of cogongrass, Imperata cylindrica, and torpedograss, Panicum repens. Weed Sci. 36: 577-582.

Wilcut, J. W., B. Truelove, D. E. Davis and J. C. Williams. 1988b. Temperature factors limiting the spread of cogongrass {Imperata cylindrica) and torpedograss. {Panicum repens). Weed Sci. 36: 49-55.

Willard, T. R. 1988. Biology, ecology, and management of cogongrass {Imperata

cylindrica (L.) Beauv.). Ph. D. dissertation, Univ. of Fla., Gainesville, pp. 1 13.

Willard, T. R., J. F. Gaffney, and D. G. Shilling. 1997. Influence of herbicide combinations and application technology on cogongrass {Imperata cylindrica) control. Weed Technol. 11: 76-80. 192

Willard, T. R., D. W. Hall, D. G. Shilling, J. A. Lewis, and W. L. Currey. 1990. Cogongrass {Imperata cylindrica) distribution on Florida highway rights-of-way. Weed Technology 4: 658-660. '

Willard, T. R. and D. G. Shilling. 1990. The influence of growth stage and mowing on competition between Paspalum notatum and Imperata cylindrica. Trop. Grass. 24: 81- 86.

Willard, T.R. and D.G. Shilling. 1990. The influence of growth stage and mowing on competition between Paspalum notatum and Imperata cylindrica. Trop. Grass. 24: 81- 86.

Willard, T.R., D.G. Shilling, J. F. Gaffney, and W. L. Currey. 1996. Mechanical and chemical control of cogongrass {Imperata cylindrica). Weed Technol. 10: 722-726.

Winder, R.S. and C.G. Van Dyke. 1987. The effect of various adjuvants in biological

control of Johnsongrass (5. halepense) with the fungus B. sorghicola. Proc. Weed Sci. Soc. 27:128.

Winder, R. S. and C. G. Van Dyke. 1990. The pathogenicity, virulence, and biological potential of two Bipolaris species on Johnsongrass {Sorghum halepense). Weed Sci. 38: 89-94.

Winston, P.W. and D. H. Bates. 1960. Saturated solutions for the control of humidity in biological research. Ecology 41(1): 232-237. ^*

Womack, J.G. and M.N. Burge. 1993. Mycoherbicide formulation and the potential for bracken control. Pestic. Sci. 37: 337-341.

Womack, J.G., G.M. Eccleston, and M.N. Burge. 1996. A vegetable oil-based invert • emulsion for mycoherbicide delivery. Biol. Control 6: 23-28.

' Wymore, L. A., C. Poirier, A. K. Watson, and A. R. Gotlieb. 1988. Colletottrichum coccodes, a potential bioherbicide for control of velvetleaf {Abutilon theophrasti). Plant Dis.72: 534-538.

Wymore, L. A. and A. K. Watson. 1989. Interaction between velvetleaf isolate of Colletotrichum coccodes and thidiazuron for velvetleaf {Abutilon theophrasti) control in - the field. Weed Sci. 37: 478-483. .

Yang, Y-K., S-O. Kim, H-S. Chung, and Y-H. Lee. 2000. Use of Colletotrichum graminicola KAOOl to control bamyardgrass. Plant Dis. 84: 55-59.

Yoder, O. C. 1976. Page 16, in: Biochemistry and Cytology of Plant Parasite

Interactions, K. Tomiyama et al, eds. Elsevier Publ. Co., Amsterdam. 193

Echinochloa species Zhang, W. M., A. K. Watson, and Moody K. 1996. Responses of Plant Dis. 80(9): 1053-1958. and rice (Oryza sativa) to indigenous pathogenic fungi.

monoceras, a Zhang, W. M., and A. K. Watson. 1997. Host range of Exserohilum J. Bot. 75: 685-692. potential bioherbicide for the control of Echinochloa species. Can.

Cereals: Guide to Zillinsky, F.J. 1983. Common Diseases of Small Grain A Trigo. Identification. Centro Intemacional de Mejoramiento de Maiz y

Perspectives on providing a realistic Zomer, P. S., S. L. Evans, and S. D. Savage. 1993. 79-86, in: Pest technical formulation for the commercialization of bioherbicides. Pages J. R. Plimmer, Control with Enhanced Environmental Safety, S. O. Duke, J. J. Menn, and eds. Washington, D. C: American Chemical Society.

Zwolfer, H. and Harris, P. 1971. Host specificity determination of insects for biological control of weeds. Ann. Rev. Entomol. 16:159-178. 194

BIOGRAPHICAL SKETCH

1969. Camilla B.Yandoc was bom in Baguio City, Philippines, on October 28,

Francisco Yandoc and Marietta Balasoto. She is the second among the four children of

majored in Plant She entered the University of the Philippines at Los Banos in 1986. She

Pathology and received a Bachelor of Science degree in Agriculture in 1991.

of She worked as a researcher at the Entomology and Plant Pathology Division

the International Rice Research Institute (IRRI) from 1991 to 1995. She attended

to graduate school at the University of the Philippines for two years before deciding

pursue graduate studies in the United States. She started on a Master's program at the

Plant Pathology Department of the University of Florida in the spring of 1996 and

eventually switched to a straight Ph.D. program. She married Gino P. Abies in

September 1998. She received the F.A. Wood Award in Plant Pathology and was

inducted into the Gamma Sigma Delta Honor Society of Agriculture University of

Florida Chapter in 2000. . . , , it to I certify that I have read this study and that in my opinion conforms acceptable standards of scholarly presentation and is fully adequate, in scope and quality as a dissertation for the degree of Doctor of Philosophy.

Raghavan Charudatfan, Chair Professor of Plant Pathology

to I certify that I have read this study and that in my opinion it conforms acceptable standards of scholarly presentation and is fully adequate, in scope and quahty as a dissertation for the degree of Doctor of Philosophy.

Richard D. Berger ^ Professor of Plant Pathology

it to I certify that I have read this study and that in my opinion conforms acceptable standards of scholarly presentation and is fully adequate, in scope and quaUty as a dissertation for the degree of Doctor of Philosophy.

James W. Kimbrou Professor of Plant Pathology

I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation andjs MLy_adequate, in scope and quality as a dissertation for the degree of Doctor of Philosophy.

Isf ^ -'^-TG. Shilling Professor of Agronomy . . .

This dissertation was submitted to the Graduate Faculty of the College of Agriculture and to the Graduate School and was accepted as partial fulfillment of the requirements for the degree of Doctor of Philosophy.

December 2001 .

. Dean, College of Agricultur; i. and Life Sciences

Dean, Graduate School