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).