II. GROWTH,BENTGRASSDEAD SPOTREACTIVATION,PSEUDOTHECIAPRODUCTIONANDASCOSPOREGERMINATIONOF OPHIOSPHAERELLA AGROSTIS

SYNOPSIS

Ophiosphaerella agrostis Dernoeden, M. P. S. Camara, N. R. O'Neill, van

Berkum et M. E. Palm is a newly described pathogen, which incites bentgrass dead spot

(BDS) of creeping bentgrass (Agrostis palustris Huds.). Little is known about the

biology of 0. agrostis, hence the primary goal of this study was to determine some basic

biological properties of the pathogen and epidemiological components of the disease.

The study objectives were: 1) to determine the cardinal temperatures for growth of 0.

agrostis and to describe colony morphology at each temperature; 2) to assess reactivation

of the disease from winter-dormant, field infected samples at various temperatures; 3) to

develop a technique to produce pseudothecia and ascospores in vitro; and 4) to evaluate

factors that promote ascospore germination. The growth rate of ten isolates of 0. agrostis

was compared at six temperatures (i.e., 10, 15,20,25,30, and 35°C). Differences in

growth among the isolates were apparent at all temperatures, but the optimum growth rate

for all isolates was either 25 or 30 DC. Previously infected, winter dormant creeping

bentgrass samples were collected between 1999 and 2001, and incubated at temperatures

ranging from 15 to 30°C. Between 12 and 28 d of incubation, BDS reactivation occurred

at temperatures ~ 20°C, but the greatest increase in BDS patch diameter occurred in

samples incubated at 25 and 30°C. Samples incubated at 15 °C did not develop disease

symptoms. The pathogen, however, was isolated from the leaves of plants incubated at

all temperatures, showing that 0. agrostis is capable of surviving in or on dormant tissue.

Pseudothecia were produced in vitro on a tall fescue (Festuca arundinacae Schreb.) seed

5

and wheat (Triticum aestivum 1.) bran mix infested with an isolate of 0. agrostis.

Pseudothecia developed under constant light on a benchtop growth hood (13-28 0c) or in

an incubator (25°C). No pseudothecia developed in darkness or on a lab bench in

ambient light. Pseudothecia developed in as few as 4 d of incubation and mature

ascospores were present after 7 d. Maximum pseudothecia production occurred between

28 and 31 days of incubation. Ascospores were observed to be forcefully ejected under

light or oozed en masse from ostioles in the presence of water. Ascospores from

pseudothecia produced in vitro, and incubated at 25°C, germinated in as little as 2 h.

Germ tubes generally emerged from the terminal rather than interior cells of the

ascospores. Germination during the first 4 h of incubation was enhanced by both light

and the presence of bentgrass leaves or roots. After 18 h of incubation, however, there

were few differences in ascospore germination percentages among light and dark

treatments or the presence or absence of plant tissue.

6

INTRODUCTION

Creeping bentgrass (Agrostis palustris Huds; synonym A. stolonifera L.) is a

commonly used turfgrass species on golf course putting greens throughout the United

States because of its ability to withstand low mowing heights and intense cultural

practices, and because it provides a high quality putting surface. In 1999, Dernoeden et

al. discovered a new disease of creeping bentgrass incited by an unidentified species of

Ophiosphaerella. Through morphological and molecular studies, it was shown that the

pathogen constituted a new species, Ophiosphaerella agrostis Dernoeden, M. P. S.

Camara, N. R. O'Neill, van Berkum et M. E. Palm (Camara et aI., 2000). The disease

was named bentgrass dead spot (BDS) (Dernoeden, 2000).

On close-mown creeping bentgrass grown on golf course putting greens, BDS

appears initially as small, reddish-brown spots approximately 1.0 cm in diameter and

increasing to about 8.0 cm in diameter (Dernoeden et al., 1999). During early stages of

disease development, the spots are reddish-brown or copper-colored and mimic ball-mark

injury. As the disease progresses, grass in the center of the spots becomes tan, while

leaves in the outer edge appear reddish-brown. Patches may be distributed throughout the

putting green or localized, but the spots and patches generally do not coalesce.

Sometimes the spots form depressions or pits in the putting surface. Spots recover very

slowly, as stolon growth into dead patches appears restrained or inhibited. Foliar

mycelium is not observed in the field, but when diseased plants are incubated under high

humidity for 3 to 5 days a white to pale pink foliar mycelium may develop. Numerous

pseudothecia can be found on necrotic leaf, sheath, and stolon tissues.

7

Spegazzini (1909) described 0. graminicola Speg., the type species of the genus,

which he found to be a pathogen of sprangletop (Leptochloa virgata (1.) P. Beauv.) in

Argentina. Three other turfgrass pathogens within the genus Ophiosphaerella have been

described. Ophiosphaerella herpotricha 1. C. Walker, 0. korrae Walker and Smith

(formerly Leptosphaeria korrae), and 0. narmari Wetzel, Hulbert and Tisserat (formerly

Leptosphaeria narmari) were found to cause spring dead spot of bermudagrass (Cynodon

dactylon (L.) Pers) (Crahay et al., 1988; Endo et al., 1985; Smith 1965; Tisserat et al.,

1989; Walker and Smith, 1972; and Wetzel et al., 1999). Ophiosphaerella herpotricha

also causes spring dead spot in buffalo grass (Buchloe dactyloides (Nutt.) Engelm)

(Tisserat et al. 1999). Necrotic ring spot of creeping red fescue (Festuca rubra var.

rubra), and Kentucky (Poa pratensis 1.) and annual (Poa annua 1.) bluegrasses is incited

by 0. korrae (Demoeden et al., 1995; Landschoot, 1996; and Worf et al., 1986). All of

the aforementioned Ophiosphaerella species, except for 0. graminicola, are turfgrass

root pathogens. The three root pathogens are all characterized as producing darkly

pigmented hyphae on roots, but none have been reported to infect creeping bentgrass.

There is little or no information regarding the biology of the pathogen or the

epidemiology of the disease. The goal of these studies was to elucidate some basic

biological aspects of the pathogen. The primary objectives of this investigation were: 1)

to determine the cardinal (maximum, minimum and optimum) temperatures for growth of

0. agrostis and to describe colony morphology at each temperature; 2) to assess

reactivation of the disease from winter-dormant, field infected samples at various

temperatures; 3) to develop a technique to produce pseudothecia and ascospores in vitro;

and 4) to evaluate factors that promote ascospore germination.

8

MATERIALS AND METHODS

Cardinal Temperature for Growth of O. agrostis

The growth rate often isolates of 0. agrostis (OpMD-I, 3, 4, 5, 8, 10, OpVA-I,

OpOH-I, OpIL-I, and OpPA-I) from the original ten golf courses diagnosed with BDS in

1998 were compared at six temperatures (i.e., 10, 15,20,25,30, and 35°C). A 5-mm

disk of mycelium and potato dextrose agar (PDA) was removed from the outer edge of an

actively growing colony and placed in the center of a petri dish containing 15 to 20 ml of

PDA. The dishes then were sealed with parafilm and incubated in the dark. Colony

diameters were measured with a ruler in two directions to obtain a mean every 24 h until

the fastest growing isolate crossed the plate. Colony growth of each isolate between day

two and day 10 was determined to be linear. Average daily colony growth, therefore, was

determined beginning at day two. The experimental design within each incubator was a

completely randomized design with four replications. The experiment was repeated in

different growth chambers. Average daily growth data were analyzed as a 6 by 10

factorial using SAS MIXED procedure and means were separated using the Tukey least

significant difference test (SAS Institute, Inc., Cary, NC, 1999).

Bentgrass Dead Spot Reactivation Study

Regrowth of the pathogen was assessed using once-active, winter dormant disease

samples from Lowe's Island Golf Course, Sterling, VA cultivar 'Pennlinks', Inniscrone

Country Club, Avondale, PA cultivars 'L-93' + 'Providence' + 'SR 1020', and the

University of Maryland Turfgrass Research Facility, College Park, MD cultivar 'L-93'.

9

The plugs from VA and MD were from the putting surface, whereas, the plugs from PA

were from the higher cut collar. Samples collected in all three years were grown on sand-

based mixes. Three applications of chlorothalonil (tetrachloroisophthalonitrile) and

iprodione (3-(3,5-dichlorophenyl)- N-(1-methylethyl)-2,4-dioxo-1-

imidazolidinecarboxamide) were made between 19 October and 15 November 1998

before the VA plugs were collected on 1 February 1999. Greens in VA were overseeded

with creeping bentgrass during the autumn of 1998. Plugs from PA also received several

applications ofiprodione and thiophanate methyl (dimethyl-4-4'-0-phenylenebis-3-

thioallophanate) between 2 October and 24 November 1999, prior to collecting plugs on

28 February 2000. The plugs from MD received applications of chlorothalonil,

triadimefon (1-(4-chlorophenoxy)-3,3-dimethly-1-(1H-1 ,2,4-triazol-1-yl)-2-butanone)

and fenarimol (a.-(2-chlorophenyl)-a.-( 4-chlorophenyl)-5-pyrimidinemethanol) between

20 October and 10 November 2000, prior to collecting plugs on 17 January 2001.

Four, 100 rom diameter by 50 rom deep plugs containing small diseased spots (14

to 48 rom) were watered to field capacity, sealed in plastic bags, and placed in growth

chambers at four temperatures (15, 20, 25, and 30 0q. In the second and third years, but

not the initial year, an additional four plugs without disease symptoms were incubated in

each growth chamber under the aforementioned conditions. All plugs were subjected to a

12 h photoperiod. Photosynthetically activated radiation (PAR) was measured from

inside the plastic bag with a Quantum Sensor (Apogee Instruments Inc., Logan, UT). The

distance between the light source and diseased samples in the four growth chambers was

adjusted to provide an average PAR of 88 !lmol mo2sec.l (range = 77 to 93 mol m,2sec,I).

10

In all three studies, diseased and symptomless samples were acclimated on a lab bench

for 24 hr prior to incubation in the growth chamber. Plugs were completely randomized

in the growth chamber and there were four replicates per temperature treatment. Plugs

received approximately 50 ml of distilled water every 3 to 5 d as the soil began to dry out.

Leaves were trimmed to approximately 1 cm in height as needed, and plugs were

monitored every 2 to 10 d for disease symptoms. Re-isolation of 0. agrostis was

attempted during and at the completion of each experiment.

Disease reactivation and development were assessed by measuring patch diameter

in two directions with a ruler and mean patch diameter was used for the statistical

analyses. Data from each year were used to determine the area under the disease progress

curVe (AUDPC). The AUDPC values were calculated using the formula: I [(Yi+

Yi+I)/2][~+1- ~], where i = 1,2,3... n-1, Yiis the mean diameter of the BDS infection center;

and ~ is the time of the i th rating (Campbell and Madden, 1990). The AUDPC data then

were standardized by dividing the AUDPC value by the total time duration (~- tl) of the

experiment (i.e., 40 to 41 d). Mean patch diameter data also were analyzed on individual

rating dates using the SAS MIXED procedure and means were separated based on the

protected (P < 0.05) least significant difference multiple mean comparison test (SAS

Institute, Inc., Cary, NC, 1999).

Pseudothecia Production

A readily available source of spores is needed to conduct ascospore germination

studies. Hence, it was necessary to be able to produce pseudothecia in the laboratory. A

mix of tall fescue (Festuca arundinacae Schreb.) seed and wheat (Triticum aestivum L.)

11

bran 50/50% v/v was evaluated as a potential pseudothecia culture medium for 0.

agrostis. The medium was prepared by soaking tall fescue seeds in tap water overnight.

Seeds then were rinsed three times and mixed with wheat bran (v/v), placed in 50 ml

flasks, and autoc1aved for 1 h on two consecutive days. Mycelia from the edge of an

actively growing colony (OpMD-IO) was removed and placed on the surface of the

cooled seed/bran mix. Flasks then were incubated in a growth chamber under constant

light (88 J..lmolm-2 sec' Ifrom four F20TI2/CW, 20 watt bulbs (Philips Lighting,

Somerset, NJ) at 25°C for 16 to 18 d. Inoculum was mixed daily to promote aeration and

to allow mycelium to become evenly distributed throughout the medium. Approximately

0.25 g of the infested mix then was placed on sterile, moist filter paper (Qualitative 415,

VWR Scientific, West Chester, PA) in a 60 by 15 mm petri dish (VWR Scientific, West

Chester, PA). Petri dishes containing infested media were place in one of three

conditions as follows: 1) on a lab bench in ambient light (i.e., approximately 12 hrs 4

J..lmolm-2 sec.1 light); 2) in a benchtop growth hood (Model # 11000 Labconco

Corporation, Kansas City, MO) under constant light (17 Ilmol m-2 see-I) supplied from a

fluorescent, cool-white, 30 watt bulb (F30T8/CW, General Electric, Fairfield, CT); or 3)

in a growth chamber (I30BLL, Percival Scientific, Inc., Perry, IA) at 25C in constant

light (72 J..lmolm-2 see-I) from a fluorescent, cool-white, 20 watt bulb (F20TI2/CW,

Philips Lighting, Somerset, Jl.!J). Temperatures on the lab bench ranged from 13 to 28°C.

Treatments also were subjected to constant dark by covering petri dishes with aluminum

foil and placing them in the three aforementioned conditions. The inoculum and filter

paper were kept moist throughout the study. The medium was monitored daily for the

development of pseudothecia.

12

Pseudothecia production was rated on a 0 to 5 scale where 0 = no pseudothecia

observed and 5 = medium covered with pseudothecia. The experiment was arranged as a

2 x 3 factorial with three replications. The study was repeated and data from each study

were combined for the statistical analysis. Data from treatments in which pseudothecia

developed were analyzed. At each rating date, pseudothecia ratings were analyzed using

the SAS MIXED procedure and means were separated based on the protected (P < 0.05)

least significant difference multiple mean comparison test (SAS Institute, Inc., Cary, NC,

1999). In addition, regression analyses were performed to determine the rate of

pseudothecia production for each treatment and regression lines were compared using the

method described by Neter and Wasserman (1974).

Ascospore Germination

Ascospores were obtained from pseudothecia produced in vitro. Approximately

0.25 g of the infested tall fescue/wheat bran mix was placed on moist filter paper in a

petri dish and placed in a growth chamber at 25°C under constant light as previously

described. Mature ascospores were present approximately one month after incubation

(i.e., forcefully discharged when placed under direct light) and an ascospore solution was

prepared by wrapping the infested mix in cheesecloth and submerging it in 100 ml

distilled water for 2 min. Pseudothecia were not surface disinfested because mature

ascospores would exude into the disinfectant. The concentration of ascospores in the

solution was determined with a Levy haemocytometer (VWR Scientific, West Chester,

PA) and dilutions were made to obtain approximately 7.5 x 102 ascospores ml-1• A 100 III

aliquot of the suspension (about 75 ascospores) was placed on a microscope slide with

13

three, 10 mm sections of creeping bentgrass leaves or roots or with no bentgrass tissue

(distilled water control). Bentgrass leaves and roots were obtained from 3-month old 'L-

93' creeping bentgrass grown in a greenhouse. To grow the plants, approximately 0.03 g

of surface disinfested (10% Clorox'li\ 0.05% sodium hypochlorite) 'L-93' creeping

bentgrass seeds were placed in fifty, 25 cm2pots filled with a sterilized sand mix (93%

sand, 2% silt and 5% clay) with a pH of 5.8 and 9.5 mg organic matter gr.1of soil. Prior

to seeding, the soil medium was autoclaved at 121C for 1 h on two consecutive days.

Seeded pots were placed on a plastic tray and grown on a greenhouse bench. Plants were

fertilized using 109 3.8 L-Iof a 20N-20P20s-20K20 fertilizer (Jack's Classic@, lR.

Peters, Inc., Allentown, PA). Micronutrients including B, Cu, Fe, Mn, Mo and Zn also

were included in each fertilizer application for a total of 0.002, 0.005, 0.01, 0.005,

0.00009 and 0.005 g 3.8 L.1, respectively. The dissolved fertilizer solution was used to

fill the tray monthly for a total of 3 applications, and nutrients were allowed to diffuse

into the soil. Leaf and root tissue were washed in running tap water for 60 min prior to

incubation with ascospores.

Slides were placed inside two, plastic humidity boxes lined with moistened paper

towels and incubated at 25°C in either constant light (88 ~mol m.2sec.l) or darkness.

Aluminum foil was used to cover the humidity box for dark treatments. Distilled water

was periodically added to each slide to prevent ascospore desiccation. Individual slides

were removed from the incubator at 2,4,6,8, 12, 18, and 24 h for observation.

Ascospores were fixed and stained in lactophenol-cotton blue (VWR Scientific, West

Chester, PA) and slides were sealed with Cytoseal 60 low viscosity mounting medium

(Stephens Scientific, Kalamazoo, MI). Germinating spores were counted and percent

14

germination was determined. For ascospores in which germination had occurred, the

number of germ tubes developed and their respective location within each ascospore were

determined. The slides were completely randomized in a growth chamber with 3

replicates. The study was repeated and percent germination data were analyzed using the

SAS MIXED procedure. Significantly different means were separated based on the

protected (P < 0.05) least significant difference multiple mean comparison test (SAS

Institute, Inc., Cary, NC, 1999).

15

RESULTS AND DISCUSSION

Cardinal Temperature for Growth of O. agrostis

Analyses of 0. agrostis daily growth data for each isolate revealed no significant

(P < 0.05) differences between the two studies. Data from both studies, therefore, were

combined for statistical analyses. The two-way temperature x isolate interaction was

significant (P < 0.0001).

Very little growth (~0.5 mm 24 hr-I) was exhibited from all isolates when

incubated at 35°C (Table 1). When incubated at the lowest temperature (i.e., 10°C), the

growth rate of all isolates was slow, ranging from 0.7 to 1.5 mm 24 hr-l. The growth of

each isolate, however, increased as temperature increased between 10 and 25°C. With

few exceptions, the maximum growth rate for the isolates occurred equally at 25 and 30

°C and ranged from 4.8 to 6.1 mm 24 hr-l and 4.1 to 5.5 mm hr-l, respectively. The

exceptions were OpMD-1, OpMD-3 and OpPA-1, which grew faster at 25°C when

compared to 30 °C. At each temperature, OpMD-1 and OpMD-1 0 generally exhibited the

greatest and slowest growth, respectively, among all isolates. When incubated at 30°C

on PDA, however, the growth rate ofOpMD-10 was similar to most other isolates.

Colony morphology and color varied at different temperatures. At lower

temperatures, colony morphology appeared as a fluffy mycelial growth, but as

temperatures increased colonies developed into a condensed mycelial growth habit.

Hence, colonies incubated at 10 to 25°C exhibited a fluffy, aerial growth, while at 30 and

35 °C mycelial growth only was slightly raised, and the mycelium grew more closely

appressed to the medium.

16

Isolates grown at 10°C exhibited white fluffy aerial mycelium. As the colony

grew, the older mycelium in the center of the colony developed an olive-green color. The

underside of the plate was dark brown in the center, but gradually developed a light tan to

beige color toward the edge of the colony.

Isolates grown at 15°C initially appeared white, turning pink by day 3 and then

light gray to olive-green in the center by day 7. The outer edge of the actively growing

colony appeared buff or off-white. The underside of each plate initially had a dark-brown

to black center, while the outer edge of the colony appeared beige or yellow in color. An

exception was OpOH-l, which remained gray to olive green in the center, while the outer

edge of the colony was white. The underside of petri dishes containing OpOH-l

generally was dark-brown to black with a small amount of beige near the extreme edge of

the actively growing colony.

Isolates grown at 20°C initially appeared rose-quartz and turned slightly gray in

the center by day 5. A buff, outer edge persisted throughout the 10 d of incubation. The

undersides of plates were generally beige or yellow in color, but darkened to a dark

brown or black color toward the center of the colony. An exception was the isolate

OpOH-l, which appeared gray to olive-green in the middle and white at the outer edge of

the colony. The underside of the OpOH-l plates appeared brown or black, except at the

edge of the colony, which was beige in color.

At 25°C, the isolates initially developed a rose-quartz color, with a buff outer

edge. After 5 to 7 d of incubation, the center of colonies generally had a gray appearance.

The isolate OpOH-l had a mix of gray and pink-colored mycelium in the center, while

most of the colony had a fluffy, white mycelium. The underside of the plate for OpOH-l

17

was a dark brown to black color in the center and beige or tan at the actively growing

outer edge of the colony. Colonies with a rose-quartz color mycelium generally

maintained a beige or tan color on the underside of the plate. As the colonies began to

exhibit a gray appearance (i.e., 5 to 7 d after incubation) the underside of the plate began

to resemble that of OpOH-l (i.e., brown to black).

At 30 °C, aerial mycelium was considerably more condensed for most isolates.

Colonies were dark pink to red, and had a buff outer edge. The underside of these

isolates varied in color from pink to beige or brown. All of the isolates, except for

OpOH-l, grew slightly appressed and had a velvety appearance on PDA. Unlike the

other isolates, OpOH-l had a fluffy, white to olive-gray colored mycelium, which

remained evident at the end of the 10 d incubation period. In addition, the underside of

the plate for OpOH-I was a dark brown to black color in the center and beige or tan

towards the edge of the colony.

With the exception ofOpV A-I and OpMD-lO, the isolates incubated at 35 °C

grew very little and exhibited a 'knotted' growth habit on PDA. Growth appeared to be

occurring only on the original transferred PDA plug, rather than on the PDA in the plate.

During the last days of incubation, the mycelium that grew on the PDA 'lifted', and

pulled-back towards the seeded plug, giving the colony a knotted appearance. The isolate

OpMD-4 maintained a pink or red color, while the colony color of all other isolates

remained gray throughout the study. Virtually no growth on PDA was observed from

OpMD-iO and OpVA-I, but both isolates developed a deep red color in the area of the

seeded plug.

18

The optimum temperature for growth of all isolates was between 25 and 30°C.

Growth and morphology of 0. agrostis, however, varied among isolates at each

temperature. This variation among isolates, particularly OpOH-1, may be attributed to

the high frequency of meiotic recombination caused by sexual rather than clonal

repr.oduction (Camara et al., 2000). Very little or no growth occurred at temperatures as

low as 10 or as high as 35°C, indicating that growth of 0. agrostis was restricted by

relatively low temperatures (i.e., 10°C) or high temperatures (i.e., > 30 0C). The

optimum temperatures for growth of 0. agrostis in vitro (i.e., 25 to 30°C), however,

exceeded temperatures that are favorable for shoot growth of cool-season grasses (i.e., 16

to 24°C) (Beard, 1973). Bentgrass dead spot generally appears in the field in the summer

when daytime ambient air temperatures exceed 25°C. Hence, 0. agrostis appears to have

a competitive growth advantage over bentgrass in the summer.

Colony morphology and color and growth rate of 0. agrostis on PDA can be used

to distinguish this pathogen from other Ophiosphaerella spp. Ophiosphaerella

herpotricha grown on PDA produces a white, cottony mycelium that turns tan or brown

within 3 to 7 d (Tisserat et aI., 1989). Wetzel et al. (1996) reported a brownish-black

liquid exuding from the center of 0. herpotricha isolates after 2 weeks of incubation on

half-strength PDA. In addition, optimum growth of 0. herpotricha occurred at 20 to 25

°C and maximum colony gro~ was between 3.5-4.1 mm 24 h-I on PDA (Tisserat et al.,

1989). On PDA, maximum colony growth for 0. korrae and 0. narmari is 4 to 5 mm 24

h-I at 25°C (Walker and Smith, 1972). When incubated at 25°C on PDA, aerial

mycelium of 0. korrae and 0. narmari initially is white to buff and darkens as colonies

age (Walker and Smith, 1972; Wetzel et al., 1996). Ophiosphaerella korrae colonies

19

have a distinctive raised or "domelike" growth habit (Wetzel et al., 1996).

Ophiosphaerella herpotricha and 0. korrae exhibit little or no growth when incubated at

30°C on PDA (Crahay et al., 1988; Tisserat et al., 1989). Growth of 0. narmari at 30°C

has not been reported. It can be presumed, however, that when incubated at 30 °C, daily

growth of O. narmari grown on PDA is < 4 rom 24 h-I based on its optimum growth rate

of 4-5 mm 24h-1 at 25°C (Walker and Smith, 1972). When incubated at 25°C, 0.

agrostis generally can be differentiated from other Ophiosphaerella spp. by the rose-

quartz colored colony. The only exception was OpOH-l, which produces an olive-green

colored mycelium. Numerous other 0. agrostis isolates (n=42) that have been collected

also produce the distinctive rose-quartz color at 25°C on PDA (Chapter III). Differences

in colony color associated with OpOH-l, however, may add to the difficulties of properly

distinguishing 0. agrostis at 25°C. However, 0. agrostis can be distinguished from

other Ophiosphaerella spp. by the generally dark pink to red, velvety mycelium and

faster growth rate (i.e., ave = 5.1; range = 4.1-5.3 rom 24 h.l) when incubated at 30°C.

Reactivation Study

Previously diseased plugs from VA, MD and PA were incubated at four

temperatures (i.e., 15,20,25, and 30°C) in 1999. There were no healthy control plugs

from the VA site. For VA plugs, there was no visual evidence of reactivation of the

disease before 28 d. After 28 d, however, the patch diameter of plugs incubated at both

25 and 30°C began to increase. By day 40, plugs incubated at 25 and 30 °C had active

disease symptoms (i.e., leaf discoloration and death) and increased patch sizes (Fig. 1).

20

Plugs incubated at 25°C exhibited the most rapid increase in patch development and had

the highest AUDPC values (Table 2).

Adventitious roots emerged at the soil surface from the stem base or crown, and

encroachment of these roots into dead spots was observed at day 5 in plugs incubated at

15 and 20°C, and at day 16 from plugs incubated at 25 and 30 °C. Although many roots

from healthy plants along the outer edge of active patches grew into the center of dead

spot zone, roots remained at the soil surface and did not move down through the soil

profile. No signs of the pathogen (i.e., hyphae) or disease symptoms were observed on

these surface roots. The site was overseeded prior to collecting plugs, and healthy

creeping bentgrass seedlings emerged in the dead spots. Survival of the seedlings was

not noted, as seedlings were removed as they germinated. All plugs needed to be

trimmed throughout the experiment, however, after 28 d turf plugs incubated at 25°C had

little foliar growth and needed only minimal trimming. Ophiosphaerella agrostis was

isolated from all four plugs maintained at 15 and 20°C, but due to death of all plants and

presumably 0. agrostis as well, the pathogen was not isolated from plugs maintained at

25 or 30°C.

Based on the development of a bronze or reddish-brown discoloration of leaves of

plants adjacent to dead spots, reactivation ofBDS from PA plugs occurred 12 to 18 d

after incubation began at 25 and 20°C, respectively (Fig. 2). Plugs incubated at 25°C

exhibited the most rapid increase in disease development and had the highest AUDPC

value (Table 2, Fig. 2). Bentgrass dead spot symptoms developed at day 23 at 30°C, but

patch size did not increase appreciably following 40 d of incubation. Although no

symptoms were observed in plugs maintained at 15°C, 0. agrostis was isolated from

21

discolored or senescent leaves of plants maintained at all temperatures. No BDS

symptoms developed on plugs collected from the disease-free plugs at any temperature.

Prior to initiation of the trial for the MD plugs, immature ascospores were

observed inside several pseudothecia, which were found overwintering in the dead spots.

Pseudothecia were not removed. After 5 d of incubation, mycelium was observed on the

outside of empty pseudothecia (i.e., ostiole completely open) in dead spots of plugs

maintained at 25°C. Twelve d after incubation, 0. agrostis was isolated from bronze or

tan colored bentgrass leaves from plugs incubated at 20, 25, and 30°C. Plugs incubated

at 15 °C exhibited no disease symptoms and isolation of the pathogen was unsuccessful at

this time. After 16 d of incubation, patch diameter of diseased plugs maintained at 25

and 30°C began to increase. By 41 days of incubation, plugs maintained at 30 °C had the

largest patch diameter, but total disease (i.e., ADOPC) was similar to plugs incubated at

25°C (Fig. 3). Patch diameter of plugs incubated at 20°C was larger than was observed

in plugs incubated at 15 °C on day 41. The ADOPC values were similar between plugs

maintained at 15 and 20°C, but these AUDPC values were significantly less (P<0.05),

when compared to those plugs incubated at 25 and 30°C (Table 2). Although no BOS

symptoms developed on plugs incubated at 15°C, 0. agrostis was isolated from

bentgrass leaves located at the periphery of BOS patches from plugs maintained at all

temperatures on day 41. No symptoms developed on the disease-free, control plugs at

any temperature.

Winter-donnant, 0. agrostis-infected creeping bentgrass incubated at 15°C did

not develop disease symptoms. The pathogen was isolated from senescent leaves of

plants adjacent to diseased spots maintained at 15 °C in each year of the study, indicating

22

that 0. agrostis is capable of surviving in winter-dormant leaf tissue. Hyphae generally

were not observed on the roots of 0. agrostis-infected plants, as reported by Dernoeden et

al. (1999). Dark-brown to black hyphal masses and runner hyphae of 0. agrostis,

however, commonly were observed near or on the nodes of creeping bentgrass stolons.

Dark brown to black hyphae only were occasionally found in the internode region and on

roots emerging from the stem base of infected plants or the nodes of stolons. Although

BDS reactivation occurred when infected bentgrass plugs were incubated at 20 °C,

disease development was slow and AUDPC values were not significantly different from

samples incubated at 15 °C.

Crahay et al. (1988) reported that 0. korrae, the causal agent of spring dead spot,

may be capable of actively invading bermudagrass (Cynodon dactylon L.) roots as soil

temperatures decline and the host begins to enter winter dormancy. Although optimum

temperature for growth of 0. korrae is 25 °C, root growth of bermuda grass is stimulated

at this temperature, and the pathogen was unable to cause significant injury (Crahay et al"

1988). Optimum temperatures for growth of creeping bentgrass shoots and roots are 15

to 24 °C and 10 to 18 °C, respectively (Beard, 1973). Creeping bentgrass growing at 15

to 20 °C, therefore, may have a competitive growth advantage over the pathogen at these

temperatures. Although the pathogen grows on PDA at 15 to 20 °C, disease development

generally was not observed on plugs at these temperatures, indicating that temperatures

favoring bentgrass growth render plants more resistant to infection. Reactivation of BDS

may occur in as little as 12 or 16 d when incubated continuously at 25 or 30 °C,

respectively. As previously noted, in vitro studies showed that 25 and 30 °C were the

optimum temperatures for growth of 0. agrostis. Because the optimum temperatures for

23

root and shoot growth of cool-season grasses are ~ 24°C, the pathogen appears to have its

greatest competitive advantage at temperatures between 25 and 30 °C. Observations from

this study revealed that overwintering pseudothecia can germinate directly to produce

mycelia. Hence, 0. agrostis can survive winter in or on infected leaf tissue as well as

overwintering pseudothecia.

Pseudothecia Production

Several preliminary experiments were conducted to find a method to produce

pseudothecia production. In one trial, after 62 d incubation in a sealed 1L flask, the tall

fescue/wheat bran mix infested with OpVA-I was placed in a paper-lined, glass bowl,

covered and incubated on a lab bench at 13 to 28°C. Immature pseudothecia were found

5 d after the infested mix was placed on a lab bench. Mature spores were observed in 20

d (6 April 2000), and ascospores were fully mature 27 d (13 April) after the mix was

removed from the growth chamber. The pseudothecia primarily were found partially

embedded in the tall fescue seeds, but several pseudothecia formed on the paper towel.

Ascospores were ejected from the ostiole of several pseudothecia when placed under

direct light. Although this method for pseudothecia development was successful in the

initial attempt (March and April, 2000), no pseudothecia developed when the study was

repeated (December 2000 and January 2001) under these same conditions.

Another preliminary experiment helped to develop a more consistent method for

producing pseudothecia on infested tall fescue/wheat bran mix. After 134 d in a sealed

lL flask, an O. agrostis-infested mix (OpOH-I) was removed and placed on moist filter

paper (Qualitative 415, VWR Scientific, West Chester, PA) or EcoSoft™ 100% Recycled

24

Natural Roll Towels (Bay West, Harrodsburg, KY) in a sterile petri dish and incubated

under constant light in a growth hood at room temperature (i.e., 18 to 32 0q. Immature

pseudothecia were found 3 d (13 Jan 2001) after the infested mix was removed from the

growth chamber and incubated under constant light. Most pseudothecia initiated

development after 4 d incubation. Ascospores were observed being physically discharged

(i.e., reached maturity) from pseudothecium ostioles when placed under an intense, direct

light (Le., 2000 to 3000 Ilmol m-2 see-I) 3 d later (17 January 2001). The pseudothecia

primarily were found partially embedded in the tall fescue seeds, but several pseudothecia

formed directly on the towels. Production of 0. agrostis pseudothecia on tall

fescue/wheat bran mix was successful in the growth hood on the five attempts between

the January and March, 2001. These results indicated that constant light as well as

moisture plays an important role in the development of pseudothecia and ascospores in

vitro.

Because the conditions of these preliminary trials were so variable, and because

the infested mix used was between 2 and 5 months old, a more controlled experiment was

conducted. In the controlled study, inoculum was 16 to 18 d old before being placed on

filter paper in petri dishes. After 31 d incubation, no pseudothecia developed on the tall

fescue/wheat bran infested mix in any treatment subj ected to constant dark or when the

mix was maintained under diurnal, ambient light. Pseudothecia, however, developed in

the growth chamber and growth hood when exposed to constant light.

Only data from treatments in which pseudothecia developed were subjected to

statistical analyses. Initial development of pseudothecia on 16 to 18 d old media began in

both treatments between 4 to 7 d after being placed in constant light. Pseudothecia

25

development occurred linearly throughout the experiment and maximum pseudothecia

ratings (i.e., 0-5 visual scale) were observed after 28 to 31 d of incubation (Fig. 4). There

were no differences between regression lines for the growth chamber and growth hood

data. Data from both treatments, therefore, were combined to form the equation P =

O.l~I(DOI) - 0.030 (R2 = 0.88), where P = the pseudothecia rating on a 0 to 5 scale and

DOl = days of incubation. There were no differences in the development of pseudothecia

in the growth chamber or growth hood on any individual rating date (Table 3).

In summary, pseudothecia of 0. agrostis readily were produced in vitro using the

tall fescue seed/wheat bran mix and maximum pseudothecia development occurred in

about 30 d on moistened filter paper maintained under constant light, but were not

produced in darkness. Hammer and Chastagner (1987) also found that 0. korrae

pseudothecia were not produced in the dark. Pseudothecia, however, developed in as few

as 4 d and viable ascospores had developed after 7 d incubation. In addition to fruiting

bodies that developed on tall fescue seeds, pseudothecia often formed directly on

moistened filter paper lining petri dishes. Ascospore (n=lOO) length (ave.=135 f.lm,

range=80-175 Ilm) and the number of septations (ave.=13, range=6-15) generally were

within the range reported by Camara et al. (2000) (data shown in Chapter III). Several

ascospores, however, were longer than the 150 f.lmlength (range=75-150 f.lm)reported by,

Camara et al. (2000).

Large numbers of 0. agrostis pseudothecia can be found in the field on dead

tissue in the summer. A preliminary study showed that pseudothecia developed on the

tall fescue seed/wheat bran mix in ambient diurnal light in March or April, but not

26

December or January. Hence, longer day lengths in the field in the summer likely are a

key factor in pseudothecia production in vivo. Small numbers of pseudo thecia of 0.

korrae,o. narmari and 0. herpotricha have been produced in vitro on dead tissues of

inoculated plants (Crahay et al., 1988; Smith 1965; Tisserat et aI., 1989; Walker and

Smith, 1972; Worf et al., 1986). Crahay et ai. (1988) induced 0. korrae pseudothecia by

incubating infected bermudagrass (Cynodon dactylon L.) roots for 6 wk in moistened

cheesecloth in test tubes. Mature 0. korrae pseudothecia also were obtained by

inoculating Scaldis hard fescue (Fescue longifolia Thuill.) seedlings that were grown on

water agar (Hammer and Chastagner, 1987). In all of the aforementioned studies, host

tissues were used for the development of pseudothecia. Because 0. agrostis pseudothecia

developed both on a lab bench (temperature range = 13-28 0c) and when incubated at 25

0C, temperature may be less important in ascocarp production than moisture and light. In

summary, 0. agrostis pseudothecia were produced on sterile tall fescue seed/wheat bran

mix that was incubated on moist filter paper in constant light. Hence, another

distinguishing characteristic of 0. agrostis is its ability to produce prodigious numbers of

pseudothecia in vitro in a short period oftime.

Ascospore Germination

Combined analyses of ascospore germination data revealed significant (P < 0.05)

differences between the two studies. Statistical analyses, therefore, was conducted

individually on data from Study I and II (Table 4). For each study, the highest levels of

significant (P < 0.05) interactions in the 2 x 3 x 7 factorial design were analyzed. In

addition, the means of any main treatment effects not included in the highest level

27

interaction also were separated based on the protected (P < 0.05) least significant

difference multiple mean comparison test.

In Study I, there was a significant (P < 0.001) three-way interaction among light,

tissue and hours incubated. Germination of ascospores in all treatments, except the

distilled water control (i.e., control) incubated in the dark, had begun after 2 h (Table 5).

There were, however, no significant differences in percent germination among leaf, root

and the control treatments after 2 h. After 4 h of incubation, differences in ascospore

germination among light treatments were apparent. Ascospores incubated under constant

light with either leaf or root tissue exhibited the greatest germination (i.e., 34 to 35%). In

the dark, the same tissue treatments exhibited significantly less ascospore germination

(Le., 18 to 21 %) at 4 h. Both control treatments incubated in either light or dark resulted

in the lowest percent germination (i.e., 6 to 7%) after 4 h. Ascospore germination in both

light and dark controls, however, increased significantly after 6 h of incubation. Similar

levels of germination occurred among leaf and root treatments incubated for 6 h in

constant dark and in the control in constant light. Although the greatest percentage of

germinating ascospores occurred when ascospores were incubated 6 h with leaves or

roots in the presence of light, there were no differences among the aforementioned

treatments and the root or control treatments incubated in darkness. With few exceptions,

ascospore germination increased in all treatments from 6 to 8 h of incubation.

Ascospores in the control treatment incubated in the dark for 8 h, however, exhibited the

least germination (i.e., 35%), which was similar to the percentages observed after 6 h

(Le., 31%). After 8 h, the greatest number of germinated ascospores (i.e., 76 to 79%)

occurred in the leaf and control treatments incubated in light; whereas, fewer ascospores

28

had germinated (i.e., 57 to 63%) when incubated in the dark with either bentgrass leaves

or roots. The percentage of ascospores germinating for all treatments incubated in the

presence of light did not increase significantly after 8 h. All dark treatments continued to

exhibit increases in ascospore germination until 18 h, after which time there was little

increase in germination. Few significant differences in ascospore germination occurred

among treatments after 24 h of incubation, and all treatments exhibited 71 to 85%

germination at this time.

In Study II, the highest level of interaction (P = 0.05) was the two-way interaction

oflight x hours of incubation (Table 4). No treatment interactions involving tissue were

found, however, the main effect oftissue was significant at P = 0.001. Ascospore

germination data for light and hours of incubation, therefore, were combined for leaves,

roots and the control for each light x hour interaction. Because the main effect of tissue

was not included in any significant interactions, tissue data were combined for both light

and dark treatments for all hours of incubation.

The greatest percent germination was observed when ascospores were incubated

in the presence ofleaves vs. roots vs. the control (Table 6). The magnitude of the

difference, however, was small (3 to 4% germination). Similar to the results in Study I,

ascospore germination was observed after 2 h of incubation. At 2, 6, 8, and 12 h of

incubation, more ascospores germinated in light (12,31,48 and 57%, respectively) when

compared to those incubated in the dark (3, 18,39 and 49%, respectively). Although

there were no differences between light and dark treatments after 4, 18 or 24 h, ascospore

germination continued to increase and after 24 h of incubation nearly all ascospores had

germinated.

29

In summary, ascospore germination can occur in as little as 2 h, and the

germination of ascospores initially was enhanced by both light and the presence of plant

tissue. During the early hours of incubation, ascospores generally germinated in larger

numbers in the presence of light and bentgrass leaves or roots. Ascospores incubated

with bentgrass leaves generally exhibited similar or greater levels of germination when

compared to ascospores incubated with roots. Ascospores incubated with either tissue

type for 2 to 12 h, however, generally resulted in a greater percent germination when

compared to ascospores incubated in water alone. After 18 h of incubation, however,

there were few percent germination differences among treatments.

On average, < 2 germ tubes per ascospore developed within the first 6 h of

incubation (Appendix C. Tables 4 and 5). The maximum number of germ tubes that

developed from an individual ascospore was four, and ascospores with ~ 3 germ tubes

accounted for an average of 6 and 11% of the total percentage of germinated ascospores

in Study I and II, respectively. Germination generally occurred at the terminal ends of

ascospores and an average of 73 and 58% of the total percentage of germinating

ascospores between 2 and 6 h of incubation occurred at one or both ends of ascospores in

Study I and II, respectively. Germination from the interior cells of ascospores occurred

on a limited basis.

Sugars, amino acids and other organic compounds are exuded by leaves and roots

(Beard, 1973). Exudates from bentgrass leaves and roots, therefore, likely encouraged

the germination of 0. agrostis ascospores. Endo and Amacher (1964) found that

creeping bentgrass leaf exudates increased the rate and percent of conidia germination in

30

Bipofaris sorokiniana (Sacc.) Shoemaker (formerly Hefminthosporium sorokinianum

Sacc.). An increase in the incidence of brown patch (Rhizoctonia so/ani Kuhn) also has

been attributed to the presence of exudates on the surfaces of bentgrass leaves (Rowell,

1951). In addition to exudates, extended leaf wetness duration caused by dew (Le.,

condensation of water forming droplets on leaf surfaces) also provides a suitable

environment for ascospore germination and germ tube development.

Ascospores of 0. agrostis were ejected through ostioles of pseudo the cia in light.

In the presence of moisture, ascospores also exuded en masse from ostioles. Hence,

ascospores can be dispersed by both water and wind. In the field, dew and leaf surface

exudates develop in a bentgrass canopy at about 20:00 hrs in the summer and leaves often

remain wet until 10:00 hrs when daytime temperatures reach ~ 25 °C. Ascospores

alighting on wet leaves in the morning can begin germinating in 2 h. In unrelated studies,

germ tubes of ascospores were observed entering stomates on bentgrass leaves.

Appressorium formation and the subsequent direct penetration of the epidermal cells of

creeping bentgrass leaves and roots also was observed. Hence, leaf surface exudates

appear to be an important factor in rapid germination and subsequent infection of leaves

by 0. agrostis during the morning hours.

31

Table 1. Average daily growth of ten Ophiosphaerella agrostis isolates incubated on potato dextrose agar at six temperatures for tendays.

Isolate designation Colony diameter t10°C 15 °C 20°C 25°C 30°C 35°C

mm24 h{1OpIL-1 1.1 nopt 2.3 m 3.2 jk 5.2 bed 5.3 bed 0.4 rsOpMD-l 1.4 no 3.0 jk1 4.5 efg 6.1 a 5.5 b 0.4 rsOpMD-10 0.7 pqr 2.4 m 3.4 IJ 4.8 def 4.9 b-e 0.1 sOpMD-3 1.0 nop 2.4 m 3.8 hi 5.3 be 4.1 gh 0.2 rsOpMD-4 1.2 nop 2.6 1m 3.8 hi 4.8 cde 5.2 bcd 0.4 rs

OpMD-S 1.4 no 3.0 jkl 4.3 fgh 5.0 bed 5.3 bcd 0.4 rs

OpMD-8 1.2 no 2.5 1m 3.4 IJ 4.8 c-f 5.1 bed 0.4 rs

OpOH-l 1.2 nop 2.8 klm 3.8 hi 5.1 bcd 5.4 b 0.4 rs

OpPA-l 1.5 n 2.8 klm 4.4 efg 5.4 b 4.8 c-f 0.5 qrs

~ OpVA-1 1.0 opq 2.7 klm 4.1 gh 5.1 bed 5.2 bed 0.2 rstv

t Data are an average of two separate trials conducted in different growth chambers.t Means followed by the same letter in rows and columns are not significantly different at P < .05 according to the Tukey least

significant difference test. Means were separated by SAS MIXED using 4 significant digits, but data shown have been truncated forsimplicity.

Table 2. Effect of temperature on the area under the disease progress curve (AUDPC) for bentgrass dead spot patches from previouslyinfected plugs obtained in winter from three sites between 1999-2001.

2001Temperature

eC)

AUDPCt2000

----------------- Disease day-l15 41bc~ 29b 20b20 39c 36b 23b25 52a 55a 29a30 48ab 31b 31a

t Previously infected plugs used to determine AUDPC values were obtained from VA, PA, and MD in 1999,2000 and 2001,respectively.

t Area under the disease progress curve values were determined after 40 d of incubation in 1999 and 41 d in both 2000 and 2001.~Means in columns followed by the same letter are not significantly different at P < 0.05 according to the protected least significant

difference multiple mean comparison test.

31

5.05.0

27

4.74.3

24

4.34.3

203.34.0

1074

HoodGrowth Chamber

Table 3. Production of Ophiosphaerella agrostis pseudothecia on a tall fescue/wheat bran medium incubated in a bench top growthhood (13-28 °C) and growth chamber (25°C) under constant light.

Study I Pseudotheciat

Days of incubation13 17

---------------- 0-5 scale0.7 t 1.3 2.3 2.7 3.30.5 1.0 1.7 2.3 4.0

Study II Pseudothecia tDays of incubation

3 7 10 14 18 23 28---------------- 0-5 scale

Hood 0 t 1.7 2.0 2.7 4.3 4.7 5.0Growth Chamber 0 1.0 1.3 2.3 4.0 4.7 5.0

t Production of pseudothecia was rated on a 0 to 5 scale where 0 = no pseudothecia and 5 = inoculum covered with pseudothecia.t Means in each column for Study I and II were not significantly different (P < 0.05) based on the protected least significant difference

multiple mean comparison test.

Table 4. Analysis of variance for Ophiosphaerella agrostis ascospore germination data at various time intervals after incubation at 25°c with 'L-93' creeping bentgrass leaves or roots in either constant light or darkness.Source of variance Study I Study II

ndf ddf F Value Pr> F ndf ddf F Value Pr> F

Hours incubated (hours) 6 84 334.55 ***t 6 84 533.32 ***Tissue type (tissue) 2 84 5.91 ** 2 84 14.7 ***Light 1 84 53.66 *** 1 84 28.44 ***Hours x tissue 12 84 4.37 *** 12 84 1.6 NSHours x light 6 84 6.36 *** 6 84 3.19 **Tissue x light 2 84 8.21 *** 2 84 1.19 NS

Hours x tissue x light 12 84 3.84 *** 12 84 0.81 NS

t Not significant (NS) or **, *** = significant at P ~ 0.01, and 0.001, respectively.

wVl

Table 5. Percent germination of Ophiosphaerella agrostis ascospores in Study I at various time intervals after incubation at 25°Cwith 'L-93' creeping bentgrass leaves or roots in either constant light or darkness.

Treatment Ascospore germinationt

Hours incubatedLight Tissue 2 h 4 h 6 h 8 h 12 h 18 h 24 h

%72c-g71 d-g85 a73 b-g83 abc78 a-e

83ab73 b-g78 a-e76 a-f83 abc68 e-h

Light Leaf I3 opqt 35 kl 42jk 76 a-f 77 a-eLight Root 8 pqr 34 kl 4 I k 70 efg 64 ghLight None 5 qr 7 qr 271mn 79 a-e 81 a-dDark Leaf 3 qr 21 mno 23 mno 57 hi 53 ijDark Root 3 qr 18nop 32 klm 63 ghi 66 fghDark None Or 6qr 31 kim 35kI 64gh

t Germination data collected from 50 ascospores in each of three replications (n= 150).t Means in columns and rows for each parameter followed by the same letter are not significantly different (P < 0.05) based on the

least significant difference multiple mean comparison test.

Table 6. Germination of Ophiosphaerella agrostis ascospores in Study II at various time intervals after incubation at 25°C with 'L-93' creeping bentgrass leaves or roots in either constant light or darkness.

Treatment Ascospore germination t

--------------%-------------50a~47b43 c

12h~3i

18g18 g31 f18 g48d3ge57 c49d85 b85 b93 a92a

t Germination data collected from 50 ascospores in each of three replications for each treatment (n = 150).t Ascospores incubated in either constant light (+) or darkness (-).~ Means in the column for tissue and for hours incubated x light followed by the same letter are not significantly different (P < 0.05)

according to the protected least significant difference multiple mean comparison test.

TissueLeafRootNone

Hours incubated Lightt2 +24 +

w 4-...J 6 +

68 +812 +1218 +1824 +24

100

60 - -.------ ----------------~--- ~-------~

70 - ---~~

-e-15 C; AUDPC = 41 bc

-A- 20 C; AUDPC = 39 c

~25 C;AUDPC =52a

-e- 30 C; AUDPC = 48 ab

40

50 - ~ -~------ --.--- ---- ----- .-

90 -

80-ElEl--...~-~El~...."0-0=-'""0~~w "0

00

'"'"~-1:1~~

30 -

o 4 8 12 16 20 24 28 32 36 40Days incubated

Figure 1. Effect of temperature on the diameter of bentgrass dead spot patches from previously diseased 'Pennlinks' creepingbentgrass plugs from Virginia over a 40 d period of incubation at four temperatures, 1999. Marked points with different letters aresignificantly different (P < 0.05) based on the protected least significant difference multiple mean comparison test.

90a

cc

a

c

a

bc bc bcbc

--- ---1,-- --b------o--b---b--

-"--- ----- ---------------" --.----a--a-15 C;AUDPC = 29 b

-b- 20 C; AUDPC = 36 b

~25 C; AUDPC = 55 a

--e-30 C; AUDPC = 31 b

80

20o 4 8 12 16 20 24 28 32 36 40

Days incubated

Figure 2. Effect of temperature on the diameter of bentgrass dead spot patches from previously diseased 'L-93 + SRI020 +Providence' creeping bentgrass plugs from Pennsylvania over a 41 d period of incubation at four temperatures, 2000. Markedpoints with different letters are significantly different (P < 0.05) based on the protected least significant difference multiple meancomparison test.

70

60 -

--ee--t 50 --~e(OS

;a-&. 40 -~"0

(OS~"0~~ 30 --a..bI)-Cl~=20

-e-15 C; AUDPC = 20 b

-A:- 20 C; AUDPC = 23 b

~25 C; AUDPC = 29 a

-e-30 C; AUDPC = 31 a

-------- -- --.----------- -.- -----.. ----- --------a:-

~----~ .-_._~_.--- ---"

c

10o 4 8 12 16 20 24 28 32 36 40

Days incubated

Figure 3. Effect of temperature on the diameter of bentgrass dead spot patches from previously diseased 'L-93' creeping bentgrassplugs from Maryland over a 41 d period of incubation at four temperatures, 2001. Marked points with different letters aresignificantly different (P < 0.05) based on the protected least significant difference multiple mean comparison test.

_ Growth Hood

32

111

28

111

24

oGrowth Chamber

III III

20

III

D

o

16

Days of incubation

Dill

111111

III-

12

-III

111111-- --8--,-4 8

2

oo

3

4

5

Figure 4. In vitro development of Ophiosphaerella agrostis pseudothecia on tall fescue/wheat bran media over 31 days ofincubation (001) under constant light in a growth hood (13-28 °C) or in a growth chamber (25°C). Pseudothecia development wasrated on a 0 to 5 scale where 0 == no pseudothecia present and 5 == media covered with pseudothecia. The regression line forpseudothecia development, combined for both treatments, is P == 0.181 (DOl) + 0.030 (R2 == 0.88).