INFORMATION TO USERS

This was produced from a copy of a document sent to us for microfilming. While the most advanced technological means to photograph and reproduce this document have been used, the quality is heavily dependent upon the quality of the material submitted.

The following explanation of techniques is provided to help you understand markings or notations which may appear on this reproduction.

1.The sign or “target” for pages apparently tacking from the document photographed is “Missing Page(s)”. If it was possible to obtain the missing page(s) or section, they are spliced into the film along with adjacent pages. This may have necessitated cutting through an image and duplicating adjacent pages to assure you of complete continuity.

2. When an image on the film is obliterated with a round black mark it is an indication that the film inspector noticed either blurred copy because of movement during exposure, or duplicate copy. Unless we meant to delete copyrighted materials that should not have been filmed, you will find a good image of the page in the adjacent frame.

3. When a map, drawing or chart, etc., is part of the material being photo­ graphed the photographer has followed a definite method in “sectioning” the material. It is customary to begin filming at the upper left hand comer of a large sheet and to continue from left to right in equal sections with small overlaps. If necessary, sectioning is continued again—beginning below the first row and continuing on until complete.

4. For any illustrations that cannot be reproduced satisfactorily by xerography, photographic prints can be purchased at additional cost and tipped into your xerographic copy. Requests can be made to our Dissertations Customer Services Department.

5. Some pages in any document may have indistinct print. In all cases we have filmed the best available copy.

University Micrdnlms International 300 N ZE6B ROAD. ANN ARBOR. Ml 48106 18 BEDFORD ROW, LONDON WC1 R 46J. ENGLAND 8100162

HAGAN, AUSTIN KENT

EPIDEMIOLOGICAL STUDIES OF MELTING-OUT OF KENTUCKY BLUEGRASS AND DEVELOPMENT OF A FUNGICIDE BIOASSAY

The Ohio State University PH.D. 1980 University Microfilms International300 N Zeeb Road. Ann Arbor, MI 48106 PLEASE NOTE: In all cases this material has been filmed 1n the best possible way from the available copy. Problems encountered with this document have been Identified here with a check mark .

1. Glossy photographs______2. Colored Illustrations______3. Photographs with dark background 4. Illustrations are poor copy 5. °r1nt shows through as there is text on both sides______of page 6. Indistinct, broken or small print on several pages

7. Tightly bound copy with print lost 1n______spine 8. Computer printout pages with indistinct print _ _ 9. Page(s) lacking when material received, and not available from school or author 10. Page(s) ______seem to be missing in numbering only as text follows 11. Poor carbon copy______12. Not original copy, several pageswith blurred ____ type 13. Appendix pages are poor copy______14. Original copy with light type______15. Curling and wrinkled pages______16. Other

University Microfilms International 300 is ZSSS SO. ANN 4R90S VIi J0106 '3131 761-4700 EPIDEMIOLOGICAL STUDIES OF MELTING-OUT OF KENTUCKY BLUEGRASS AND

DEVELOPMENT OF A FUNGICIDE BIOASSAY

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the

Degree Doctor of Philosophy in the Graduate

School of The Ohio State University

By

Austin Kent Hagan, B.S., M.S.

A A A A A

The Ohio State University

1980

Reading Committee: Approved By

Dr, P. 0. Larsen

Dr. L. H. Rhodes II Aoviser Dr. R. E. Hite Department of Plant Pathology ACKNOWLEDGEMENTS

Special thanks to Dr. P. 0. Larsen, Dr. L. H. Rhodes, and Dr.

R. E. Hite for their suggestions and contribution to this project.

Research was supported in part by the ChemLawn Corporation of

Columbus, Ohio and an Ohio Turfgrass Foundation Grant.

ii VITA

August 23, 1953...... Bom, Philadelphia, Pennsylvania

1975 B.S., Indiana University of Pennsylvania, Indiana, Pennsylvania

1978 M.S., Department of Plant Pathology, The Ohio State University, Columbus, Ohio

FIELD OF STUDY

Major Field: Plant Pathology

Research Interests:

Diseases of turfgrasses.

Taxonomy and nomenclature of Drechslera, Bipolaris, and Exserohilum spp.

ill TABLE OF CONTENTS PAGE

ACKNOWLEDGEMENTS ...... ii

VITA ...... ill

LIST OF T A B L E S ...... vii

LIST OF FIGURES...... viii

LITERATURES REVIEW ...... 1

Etiology and Epidemiology of Melting-Out...... 1 Conidium Discharge...... 6 Bioassays of Fungicide Residue Activity ...... 8

Chapter

I. MELTING-OUT OF KENTUCKY BLUEGRASS: INOCULUM SOURCE, PRODUCTION, RELEASE AND DISPERSAL .... 18

ABSTRACT ...... 19 INTRODUCTION...... 20 MATERIAL AND M E T H O D S ...... 22 Conidium trapping ...... 22 Meterologlcal equipment ...... 23 Inoculum source ...... 23 Disease incidence ...... 25

RESULTS...... 25 Inoculum source ...... 25 Factors influencing the sporulation of Drechslera poae in the field...... 30 Factors influencing the release of Drechslera poae conidia ...... 35 Diurnal release of Drechslera poae c o n i d i a ...... 42 Seasonal distribution of airborne Drechslera poae conidia ...... 42 Disease incidence ...... 51

DISCUSSION...... 62 LITERATURE CITED ...... 70

II. INFLUENCE OF TEMPERATURE ON DRECHSLERA POAE ACTIVITY AND LESION DEVELOPMENT ON KENTUCKY BLUEGRASS L E A V E S ...... 73 PAGE

ABSTRACT...... 74 INTRODUCTION...... 7 4 MATERIALS AND METHODS ...... 77 Plant cultivation...... 77 I noculum ...... 77 Inoculation...... 77 Influence of temperature on pre- penetration events ...... 78 Disease severity ...... 78 Leaf lesion enlargement...... 79 Penetration and colonization of leaf tissue...... 79 Infection efficiency ...... 80 Sporulation of Drechslera poae on excised Kentucky bluegrass leaves. . . 80 Comparison of Drechslera poae sporulation on intact plants and excised Kentucky bluegrass leaves...... Influence of temperature on conidium and conidiophore mor­ phology...... 81

RESULTS ...... 82 Influence of temperature on conidium germination, germ tube elongation, and appressorium formation of Drechslera poae on Kentucky blue­ grass...... Influence of temperatures on pene­ tration and colonization of Ken­ tucky bluegrass leaves ...... 89 Influence of temperature on the development of the leaf spot phase of melting-out...... 91 Influence of temperature on infection efficiency...... 105 Sporulation of Drechslera poae on Kentucky bluegrass leaves...... 106 Influence of temperature on the morphological characters of Drechslera poae conidia and conidiophores...... 106

DISCUSSION...... L16 LITERATURE CITED...... 124

v PAGE

III. COLLODION TECHNIQUE: BIOASSAY OF FUNGICIDE RESIDUE PERSISTANCE ON KENTUCKY BLUEGRASS LEAVES...... 126

ABSTRACT ...... 127 INTRODUCTION ...... 127 MATERIALS AND METHODS...... 129 Fungicide application ...... 129 Inoculation...... 129 Meterological data...... 130

RESULTS ...... 130 DISCUSSION...... 140 LITERATURE CITED ...... 142

BIBLIOGRAPHY...... 144

vi LIST OF TABLES

TABLE PAGE

2.1 Influence of temperature on the penetration of Kentucky bluegrass leaves by Drechslera poae...... 90

2.2 Sporulation of Drechslera poae on diseased excised leaves and intact 'Park' Kentucky bluegrass plants after six days incubation in a moist environment at three temperatures...... 109

2.3 Influence of temperature on the morphological characters of Drechslera poae conidia and conidiophores developing on lesions on 'Park* Kentucky bluegrass leaves ...... 114

3.1 Correlation between time and pre-penetration events of Drechslera poae on Kentucky blue­ grass leaves treated with fungicides...... 139

vii LIST OF FIGURES

FIGURE PAGE

1.1 Populations of Drechslera poae conidia in Kentucky bluegrass thatch from March 26 to July 16, 1979...... ^6

1.2 Populations of Drechslera poae conidia in Kentucky bluegrass thatch and leaf litter from July 25 to November 2, 1979...... 2®

1.3 Populations of Drechslera poae conidia in Kentucky bluegrass thatch and leaf litter from March 17 to July 10, 1980......

1.4 Summary of weekly total rainfall and mean thatch temperature from April 4 to July 10, 1980 at the ChemLavn Research Facility, Milford Center, O h i o ...... „ ...... 33

1.5 Influence of weather variables on the pro­ duction and release of Drechslera poae conidia within a Kentucky bluegrass field from June 11-13, 1980 ...... 36

1.6 Influence of weather variables on the pro­ duction and release of Drechslera poae conidia within a field of Kentucky bluegrass from May 20- 22, 1980 ...... 38

1.7 Influence of weather variables on the production and release of Drechslera poae conidia within a Kentucky bluegrass field from May 12-14, 1980...... 40

1.8 Influence of weather variables on the production and release of Drechslera poae conidia within a Kentucky bluegrass field from June 13-15, 1979 ...... 43

1.9 Influence of weather variables on the production and release of Drechslera poae conidia within a Kentucky bluegrass field from May 15-17, 1979...... 45

1.10 Diurnal periodicity of airborne Drechslera poae conidia collected within a Kentucky bluegrass field with a Kramer-Collins spore trap from April 17 to November 2, 1979...... 47

viii FIGURE PAGE

1.11 Diurnal periodicity of airborne Drechslera poae conidia collected within a Kentucky bluegrass field with a Burkard spore trap from April 1 to July 6, 1980......

1.12 Seasonal distribution of airborne Drechslera poae conidia collected within a field of Kentucky bluegrass with a Kratner-Collins spore trap from April 22 to July 29, 1979...... 52

1.13 Seasonal distribution of airborne Drechslera poae conidia collected within a field of Kentucky bluegrass with a Kramer-Collins spore trap and Burkard spore trap from July 30 to November 2, 1979...... 54

1.14 Seasonal distribution of airborne Drechslera poae conidia collected within a field of Kentucky bluegrass with a Burkard spore trap from March 31 to July 14, 1980...... 56

1.15 Incidence of the leaf spot and crown rot phases of melting-out of Kentucky bluegrass caused by Drechslera poae in 1979...... 58

1*16 Incidence of the leaf spot and crown rot phases of melting-out of Kentucky bluegrass caused by Drechslera poae from March 24 to July 10, 1980...... 60

2.1 Influence of temperature on the germination of Drechslera poae conidia on leaves of 'Park' Kentucky bluegrass ...... 83

2.2 Influence of temperature on the elongation of Drechslera poae germ tubes on leaves of 'Park' Kentucky bluegrass...... 85

2.3 Influence of temperature on the formation of appressoria of Drechslera poae on leaves of ’Park' Kentucky bluegrass...... 87

2.4 Pre-penetration events of Drechslera poae on leaves of Kentucky bluegrass ...... 91

2.5 Penetration and colonization of Kentucky bluegrass leaves by Drechslera poae ...... 95

2.6 Influence of temperature and moisture on the incidence of the leaf spot phase of melting- out of kentucky bluegrass caused by Drechslera poae...... 97 ix FIGURE PAGE

2.7 Influence of temperature and moisture on the severity of the leaf spot phase of melting-out of Kentucky bluegrass caused by Drechslera poae...... 99

2.8 Influence of temperature on leaf lesion enlargement in a moist environment...... 101

2.9 Influence of temperature on the infection efficiency of Drechslera poae on leaves of ’Park' Kentucky bluegrass ...... 103

2.10 Sporulation of Drechslera poae on excised, diseased 'Park' Kentucky bluegrass leaves at five temperatures...... 107

2.11 Profuse sporulation of Drechslera poae on a diseased, excised Kentucky bluegrass leaf ...... 11Q

2.12 Influence of temperature on Drechslera poae conidium morphology...... 112

2.13 Influence of temperature on Drechslera poae conidium morphology...... 116

3.1 Maximum and minimum daily turf microclimate temperatures and daily rainfall from October 3-24, 1979 at the OSU Turf Plots, Columbus, Ohio...... 131

3.2 Effect of fungicide residues on Kentucky bluegrass leaves on Ih poae conidium germina­ tion ...... 133

3.3 Effect of fungicide residues on Kentucky bluegrass Reaves on D. poae on germ tube elongation * ...... 135

3.4 Effect of -fungicide residues on Kentucky bluegrass leaves on D. poae appressorium formation ...... 137

x LITERATURE REVIEW

Etiology and Epidemiology of Melting-Out

Melting-out caused by Drechslera poae (Baudys) Shoem. (-HelmIn-

thosporium poae Baudys, Helminthosporium vagans Drechs., Drechslera vagans (Drechs.) Shoem.) has been recognized as one of the most

important diseases of Kentucky bluegrass (Poa pratensis L.) in the

United States (10,11,12,13,15,24,31,48,59). Melting-out can

cause severe problems on Kentucky bluegrass throughout the growing

seasons (10). Two distinct phases of melting out, the leaf spot and

crown rot phase, were recognized by Drechsler (12,13). Generally,

the leaf spot phase is most prominent during the cool, moist periods

in the spring and fall while the crown rot phase predominates during

the warm summer months (10,11). However, the severe if sporadic in­

cidence of the crown rot phase has been reported occurring from late

April to mid-June rather than later in the summer (24,58). In addition,

poae may also cause severe infections of roots and panicles of

Kentucky bluegrass (6,20).

In Minnesota, Drechslera dictyoides has been reported as the most

important pathogen of Kentucky bluegrass (5,6,18,19,63). However,

Morrison (58) failed to isolate D_. dictyoides from field collections

but frequently isolated D. poae. Morrison (58) concluded that I), poae was mlsidentifled as D. dictyoides as a result of misinterpretation

of conidium morphology at low temperatures (7-10 C). Therefore 2 reference to the above mentioned reports ( 5, 6 , 18, 19, 63) will be assumed to relate to I), poae and not D. dictyoides.

Leaf lesions Initially appear as small water-soaked spots which enlarge into purple-brown ovals with tan centers (10,11,13). Lesions developing on the crowns of Kentucky bluegrass plants are often diffuse brown discolorations (10,11). Substantial reductions in leaf density in a Kentucky bluegrass stand are the result of crown lesion girdling the leaf sheaths (10,11).

Bipolaris sorokiniana (Sacc. ex. Sorok.) Shoem. (=Helminthospor- ium sorokinlanum Sacc. ex. Sorok.) causes leaf spot lesions on Kentucky bluegrass which closely resemble those produced by D. poae. Typically,

D. poae is most frequently collected from infected plant tissue during the spring and fall while B_. sorokiniana is almost exclusively Isolated during the summer (5,24,58,59). Drechslera poae is more common in the Northern states east of the Mississippi which receive ample amounts of rainfall throughout the growing season while 1J. sorokiniana is the predominant pathogen of Kentucky bluegrass in the western half of the country where drought stress is a severe problem (5,12,24,30,57,58,59,66).

Drechslera poae overwintered'in lesions on host tissue as well as plant debris in the form of vegetative hyphae or conidia (11,31). No perfect state of I), poae has been Identified, but immature ascocarps have been found in culture (11,22,31). Conidium production of I), poae occurred in the spring and fall whenthe weather is cool and moist (11, 24).

Couch (10) reported that D. poae conidia were carried to the leaf surface by splashing water. However, other factors could be involved in the release and dispersal of D. poae conidia. Meredith (54) using a 3

Hirst spore trap situated 1 m above Bermudagrass (Cynodon dactylon)

infested with Drechslera gigantea (Heald & Wolf) Ito (-Helminthos- porium gjganteum Heald & Wolf) noted that the incidence of airborne

conidia occurred approximately at 1000 hours. The incidence of air­ borne conidia was also highest on days following rainfall or very humid nights (54). Rapid infection of leaf and crown tissue was also

favored by cool moist weather in the spring and fall (11,24,58).

Poor sporulation of D_. poae on infested leaves collected during

the summer indicates that little inoculum was available during the

summer to initiate new infections (24).

Germination of th poae conidia was rapid on excised Kentucky bluegrass leaves at 24 to 30 C (59). Horsfall (31) observed that

at temperatures between 3-35 C, 98% of the D. poae conidia on agar had germinated within 24 hours. At 24 C, germ tube development of

poae was nearly completed 12 hours after inoculation and only

small increases in length were noted (59). Well-differentiated

appressoria developed on the tips of approximately 55% of the germ

tubes on Kentucky bluegrass leaves within 18 hours of inoculation (59).

After 18-24 hours, penetration and primary hyphal development was noted

within leaves of common and 'Merlon' Kentucky bluegrass (59). Pene­

tration of leaf tissue occurred most frequently at the junctures be­

tween epidermal or guard cells but was rare through open stomates

(59). Primary hyphae which developed at the site of penetration in

the epidermal cell or intercellular space consisted of greatly ex­

panded, thin walled, hyaline and regularly septate filaments (59). \

Both primary and secondary hyphae were usually restricted to the area of

a developing lesion although hyphae may be found in healthy tissue 4 immediately after penetration (59). No information was availa­ ble concerning possible toxin production by I), poae in host tissue.

In a moist environment, sporulation of D. poae began within five days of inoculation on excised Kentucky bluegrass leaves (59).

Halisky and Funk (24) have reported that D, poae sporulation occurs less frequently on naturally infected leaves collected from May to

October. The poor sporulation of D. poae on leaf tissue collected during the warmer months of the year was attributed to the presence of a heat induced metabolite in the plant tissue (24). Colbaugh and Beard (8) reported that Helminthosporium spp. seldom sporulated on infected plant tissue remaining attached to the plant.

Difficulties have been encountered inducing abundant sporulation of I). poae on artificial media (10,13,24,31,49,59). Several authors observed that poor sporulation of D. poae was due to an antisporula- tion factor in the media (24,49). Subsequent work has shown that abundant sporulation of D. poae can be induced on artificial media or excised leaf tissue by incubating material at 15-18 C with 12-14 hours of fluorescent illumination followed by 10-12 hours of dark­ ness (22,58,63).

Temperature has a substantial effect on conidium development

(58,63). Morrison (58) has illustrated that _D. poae conidia formed at

8-10 C were substantially longer and had twice as many septa as conidia formed at 18-20 C. As Incubation temperatures increased, conidium length and number of septatlons continued to decrease (58,63).

At temperatures above 21 C, conidiophore development also became atypical with the production of enlarged cells or germ tubes at the conidiophore apex (59,63). Turf management practices have an influence on the development and severity of melting-out (11). Excessively close clipping of

Kentucky bluegrass and application of high rates of nitrogen rich fertilizers have been linked to increases in the severity of melting- out (11,13,19,24,48,51). Colbaugh and Beard (8) have observed sub­ stantial increases in the population of Helminthosporium spp. conidia on bermudagrass turf maintained at low cutting heights with high fertility levels than on turf maintained at low cutting heights with low fertility levels. However, Gibbs at al (19) have noted that disease development was greater at higher cutting heights than at the lower cutting heights. Conversely, the application of nitrogen to diseased turf has been recommended as a means of offsetting some disease damage (11,16,48,57).

Moisture may also be an important factor influencing the devel­ opment of melting-out and similar turf diseases (11). Irrigation of 'Merlon' Kentucky bluegrass seed fields increases the severity of foliar symptoms as well as the percentage of seeds Infested with

D. poae (20), Colbaugh and Endo (9) using a mineral oil flotation technique developed by Ledingham and Chinn (47), have observed sub­ stantial increases in Bipolarls sorokiniana conidium production in leaf litter collected from drought-stressed Kentucky bluegrass turf than from Irrigated turf. Increases in conidium production was attributed to increases in the saprophytic activity of _B. sorokiniana caused by substantial releases of nutrients from remoistened leaf litter (9).

Lukens (50) observed that the incidence of melting-out increases proportionally to the amount of shade over Kentucky bluegrass turf.

The enchancement of melting-out was correlated with lower carbohydrate levels In host tissue as a result of shading or nitrogen fertilization

(51). Later work indicated that the severity of Helminthosporlum - incited diseases was not linked to carbohydrate levels in host tissue

(17,19).

Conidium Discharge

A number of techniques and instruments have been developed to collect fungal spores, pollen grains, and other material from air

(21). Basically, sampling techniques can be broken down into two categories: gravity sedimentation and inertial methods (21). Sampling techniques involving sedimentation rely on the collection of airborne conidia on horizontal surfaces such as a slide or petri dish which may contain nutrient media or other material (21,28). A number of inertial traps, which operate by drawing air through a jet or tube separating material by centrification or impaction of material on rotating surface have been developed (28). The most popular and useful traps for epidemiological studies are automatic volumetric traps (17,28,37,38).

Commercially available volumetric spore traps sample either continuously or intermittantly (27,37,38). Automatic volumetric spore traps are useful for studying the diurnal periodicity of conidium release ex­ hibited by a number of dry-spore fungi like Drechslera spp. (27).

Airborne conidia of Drechslera, Bipolaris, and Exserohilum gen­ erally exhibit a diurnal or circadian periodicity (3,46,54,56). Peak field concentration of D. gigantea, EL turcica, and B_. maydis airborne conidia occurs sometime between 0900 and 1100 hours (3,46,54,56).

The actual number of airborne conidia of different species trapped will vary considerably during an epiphytotic (46,54,65). The low

Incidence of I). gigantea and _B. maydis conidia may be due to poor sporulation on host tissue or inability of conidia to remain sus­ pended in air (54,65).

The influence of weather variables on the diurnal periodicity of Drechslera, Bipolaris, or Exserohilum conidium release from coni- diophores has been hotly debated (2,48). The debate has centered on whether conidium release is passive or violent (2,4,39,41,43,44,

46,53,54,55,64). Aylor and Day (2) contend that violent release of Helminthosporium spp. conidia is epidemiologically unimportant because the percentage of conidia focibly released was often quite low (35,53,54). Leach (40) has responded by questioning whether certain important environmental conditions were monitored by Aylor and Day (2). Leach (40) does not discount the epidemiological impor­ tance of wind, but notes that wind is just one mode of conidium release which includes rainfall and violent conidium release. The relative importance of each release mechanism changes as weather variables change (40).

Passive release of conidia depends solely on wind velocity and turbulance (1,2). Experiments in wind tunnels or enclosures indicate that puffs of wind of 3-6 m/sec were required to dislodge Bipolaris maydis conidia from conidiophores (4,64). Yet, in the canopy of corn, winds with a mean speed of 1 m/sec liberated approximately 60-75% of B.. maydis conidia from c o m leaves (3). Closer examination of this phenomenon has revealed that conidium release occurred when puffs of wind reached speeds of 5 m/sec for several microseconds although the mean wind speed was approximately 2 m/sec,(4).

Violent release of conidia is triggered by either drastic changes in relative humidity or infrared radiation (39,43,44,53,54). Meredith 8 (54,55) microscopically observed the release of I), gigantea or _E.

turcica conidia after rapid desiccation. Forcible conidium release

was attributed to mechanical stress as well as production of gases within

the conidium and conidiophore after desiccation (53,54,55). Kenneth

(35) did not observe violent conidium release of several Drechslera

spp. including p.. poae after rapid conidium and conidiophore desicca­

tion. Diurnal changes in relative humidity have been linked to the

release of E. turcica (46,56). In the laboratory, Leach (39,42,43,44)

has illustrated that drastic changes in relative humidity as well as

infrared radiation influences the violent release of _E. turcica

and p. maydis conidia. Further investigation has attributed violent

release of _E. turcica conidia to the repulsion of unipolarly charged

surface of the same polarity (41,45).

Rainfall may also influence the release of dry-spore fungi coni­

dia from plant surfaces (32,39). Jarvis (34) noted massive releases

of Botrytis cinerea conidia during heavy showers on calm nights.

Conidium release was attributed to percussion waves from impacting

raindrops on moldy fruit (29,32). Increases in the number of E. tur­ cica conidia trapped following light showers were noted by Leach (46).

Leach (40,46) suggests that release may Involve electrostatic forces

as well as mechanical forces.

Bioassays of Fungicide Residue Activity

Fungicide residues on plant foliage are constantly reduced due

to weathering caused by wind, rain, temperature and light (14) .

Fungicide retention studies on leaf surfaces were usually done in

the greenhouse with artificial rainfall serving as the primary weather­

ing agent (7,26,62). Chemical tests or symptom expression were used to evaluate fungicide retention rather than the direct effects of the residues on pathogen activity (7,52). Correlations between greenhouse tests failed to account for several weather variables important in field situations (62).

Several bioassays have been developed to evaluate the persis­ tence of fungicide residues on plant foliage in the field (36,60),

Neely (60) modified the cellophane transfer technique developed by

Neely and Himelick (61) to study the persistence of commercial fungi­ cides on the leaves of 12 woody plants. Leaf disks were collected from treated plants at weekly intervals after fungicide application and placed in Coors porcelain spot plates (60,61). A conidium sus­ pension of Monilinia fructicola was applied to a cellophane disk placed over a leaf disk (60). Residue activity was evaluated by microscopically examining each cellophane disk for germinated M. fructicola conidia (60). Ko et al (36) applied fungicides to the foliage of passion fruit, collected treated leaves at weekly inter­ vals, and inoculated the excised leaves with Altemaria altemata conidia. After 24 hours, fungicide retention was evaluated by determining the number of germinated conidia using a vertical illumina tlon microscope (36).

The collodion technique which has not yet been used to evaluate fungicide residue efficacy has several possible advantages over the two bioassays already described (23). Like the preceeding tests, pathogens applied to treated host tissue are used to evaluate fungi­ cide efficacy (23). However, fungal structures are embedded in a collodion film which is stained and mounted for later observation rather than requiring immediate attention (23,36,60). This technique permits not only the monitoring of conidium germination but also germ tube elongation and appressorium formation, which are important measures of fungicide efficacy in some instances. Hagan and Larsen

(23) have illustrated that germ tube development and appressorium formation in addition to conidium germination were critical for evaluating the efficacy of several turf fungicides. Iprodione and chlorothalonil, which provided excellent disease control did not inhibit Bipolaris sorokiniana conidium germination on Kentucky blue­ grass leaves (23). However, both materials did inhibit germ tube development and suppressed appressorium formation (23). LITERATURE CITED

1. AYLOR, D. E. 1975. Resuspension of particles from plant surfaces by wind. Pages 791-812 in: The Symposium on the Atmosphere- Surface Exchange of Particles and Gases; Richland, Washington. U.S. Atomic Energy Commission Doc. No. CONF-740921.

2. AYLOR, D. E., and P. R. DAY. 1976. Conidium release in Helmin- thosporium (letter to the editor). Phytopathology. 66:537.

3. AYLOR, D. E., and R. L. LUKENS. 1974. Liberation of Helminthos- porium maydis spores by wind In the field. Phytopathology. 64:1136-1138.

4. AYLOR, D. E., and J. Y. PARLANGE. 1975. Ventilation required to entrain small particles from leaves. Plant Physiol. 56: 97-99.

5. BEAN, G. A., and R. D. WILCOXSON. 1964, Helminthosporium leaf spot of bluegrass. Phytopathology. 54:1065-1070.

6. BEAN, G. A., and R. D. WILCOXSON. 1964. Pathogenicity of three species of Helminthosporium on roots of bluegrass. Phytopathology. 54:1084-1085.

7. BURCHFIELD, H. P., and A. GOENAGA. 1957. Equipment for producing simulated rain for measuring the tenacity of spray deposits on foliage. Contrib. Boyce Thompson Inst. 19:133-140.

8 . COLBAUGH, P. F., and J. B. BEARD. 1980. Influence of management practices on conidial production by Helminthosporium spp. on improved bermudagrass. In: P. 0. Larsen and B. Joyner, eds. Advances In Turfgrass Pathology. Harvest Publications, Cleve­ land, Ohio (in press).

9. COLBAUGH, P. F., and R. M. END0. 1974. Drought stress: an Important factor stimulating the development of Helminthos­ porium sativum on Kentucky bluegrass. Pages 328-333 in: E. C. Roberts, ed., Proc. 2nd Internal. Turfgrass Res. Conf. Amer. Soc. Agronomy.

10. COUCH, H. B. 1957. Melting-out of Kentucky bluegrass, its cause and control. Golf Course Reptr. 25:5-7.

11 12

11. COUCH, H. B. 1973. Diseases of turfgrasses. Krieger Publishing Co. Huntington, NY. 348p.

12. DRECHSLER, C. 1922. A new leaf spot of Kentucky bluegrass caused by an undescribed species of Helminthosporium. (Abstr.). Phytopathology. 13:35.

13. DRECHSLER, C. 1930. Leaf spot and foot rot of Kentucky bluegrass caused by Helminthosporium vagans. J. Agr. Res. 40:447-451.

14. EBLING, W. 1963. Analysis of the basic processes involved in the deposition, degradation, persistance, and effectiveness of pesticides. Residue Rev. 3:35-163.

15. ELLIOT, R. S. 1962. Disease damage on forage grasses. Phyto­ pathology. 52:448-451.

16. ELLIOT, R. S. 1962. The effect of soil fertility on the develop­ ment of Kentucky bluegrass diseases. (Abstr.). Phytopathology. 52:1218.

17. EVERSMEYER, M. G., C. L. KRAMER, and T. I. COLLINS. 1976. Three suction-type spore samplers compared. Phytopathology. 66: 62-64.

18. GIBBS, A. F., and R. D. WILCOXSON. 1972. Effect of sugar con­ tent of Poa pratensis on Helminthosporium leaf spot. Physiol. Plant Pathology. 2:279-287.

19. GIBBS, A. F., and R. D. WILCOXSON. 1973. The effect of nitrogen fertilization and mowing on Helminthosporium leaf spot and sugar content of bluegrass leaves. Plant Dis. Reptr. 57:544-548.

20. GRAY, P.M., and J. W. GUTHRIE. 1977. The influence of sprinkler irrigation and postharvest residue removal practices on the seedborne population of Drechslera poae on Poa pratensis 'Merlon' Plant Dis. Reptr. 61:90-93.

21. GREGORY, P. H. 1961. The microbiology of the atmosphere. Leonard Hill, London. 251p.

22. HAGAN, A. K. 1980. Isolation, Identification, and Taxonomy of Drechslera and Bipolaris species on turfgrasses. In P. 0. Larsen and B. Joyner, eds. Advances in Turfgrass Pathology. Harvest Publications, Cleveland, Ohio (in press).

23. HAGAN, A. K. and P. 0. LARSEN. 1979. Effect of fungicides on conidium germination, germ tube elongation, and appressorium formation by Bipolaris sorokiniana on Kentucky bluegrass leaves and disease incidence. PLant Dis. Reptr. 63:474-478. 13 24. HALISKY, P. M . , and C. R. FUNK. 1966. Environmental factors affecting growth and sporulation of Helminthosporium vagans and its pathogenicity to Poa pratensis. Phytopathology. 56:1294-1296.

25. HALISKY, P. M., C. R. FUNK. 1966. Environmental factors affect­ ing growth and sporulation of Helminthsporium vagans influ­ enced by management practices. Plant Dis. Reptr. 50:703-706.

26. HAMILTON, J.M. 1959. Evaluation of fungicides in the greenhouse. Pages 253-257 in: C. S. Holton et al, eds., Plant Pathology: problems and progress, 1908-1958. University of Wisconsin Press, Madison. 588p.

27. HIRST, J. M. 1952. An automatic volumetric spore trap. Ann. Appl. Biol. 39:257-265.

28. HIRST, J.M. 1959. Spore liberation and dispersal. Pages 529- 538 in C. S. Holton _et al_, eds. Plant Pathology: problems and progress, 1908-1958. University of Wisconsin Press, Madison Press, Madison. 588p.

29. HIRST, J. M . , and 0. J. STEDMAN. 1963. Dry liberation of fungus spores by raindrops. J. gen. Microbiol. 33:335-344.

30. HODGES, C. F., and W. M. BLAINE. 1974. Midsummer occurrence of Helminthosporium sorokinianum leaf spot on selected cultivars of Poa pratensis. Plant Dis. Reptr. 58:448-450.

31. HORSFALL, J. G. 1930. A study of meadow crop diseases in New York. Cornell Univ. Agr, Expt. Sta. Mem, 130, 139p.

32. INGOLD, C. T. 1970. Fungal spores, their liberation and dis­ persal. Clarendon Press, Oxford, England. 302p.

33. INGOLD, C. T. 1971. Liberation mechanisms of fungi. Pages 102- 115 in: P. H. Gregory, and J. L. Monteith, eds. Airborne Microbes, 17th Symposium of the Society for General Micro­ biology. University Press, Cambridge, England. 385p.

34. JAMES, W, R. 1962. The dispersal of spores of Botrytis clnerea Fr. in a raspberry plantation. Trans. Brit, mycol. Soc. 45: 549-559.

35. KENNETH, R. 1964. Conidial release in Helminthospora. Nature 202: 1025-1026.

36. K0, W. H., H. LIN, and R. K. KUNIMOTO. 1975. A simple method for determining efficacy and weatherability of fungicides on foliage. Phytopathology. 65:1023-1025.

37. KRAMER, C. L., M. G. EVERSMEYER, and T. I. COLLINS. 1976. A new 7-day spore sampler. Phytopathology. 66:60-61. 14

38. KRAMER, C. L., and S. M. PADY. 1966, A new 24 hour spore sampler. Phytopathology. 56:517-520.

39. LEACH, C. M, 1975. Influence of relative humidity and red-infra­ red radiation on violent spore release by Drechslera turcica and other fungi. Phytopathology. 65:1303-1312.

40. LEACH, C. M. 1976. Further comments on spore release by Drechslera (Helminthosporium) spp. and other fungi, (letter to editor) Phytopathology. 66:1265-1266.

41. LEACH, C. M. 1976, An electrostatic theory to explain violent spore liberation by Drechslera turcica and other fungi. Mycologia. 68:63-86.

42. LEACH, C. M. 1980. An apparatus for the precise control of humidity, temperature, airflow, and light in spore discharge studies. Phytopathology. 70:189-191.

43. LEACH, C. M. 1980. Influence of humidity and red-infrared radia­ tion on spore discharge by Drechslera turcica-additional evidence. Phytopathology. 70:1920196.

44. LEACH, C. M. 1980. Vibrational release of conidia by Drechslera maydis and D_. turcica related to humidity and red-infrared radiation. Phytopathology. 70:196-200.

45. LEACH, C. M. 1980. Evidence for an electrostatic mechanism in spore discharge by Drechslera turcica. Phytopathology. 70: 206-213.

46. LEACH, C. M., R. A. FULLERTON, and K. YOUNG. 1977. Northern leaf Slight of maize in New Zealand: release and dispersal of conidia of Drechslera turcica. Phytopathology. 67:380- 387.

47. LEDINGHAM, R. J., and S. H. F. CHINN. 1955. A flotation method for obtaining spores of Helminthosporium sativum from soil, from soil. Can. J. Bot. 33:298-303.

48. LEFEBVRE, C. L., F. L. HOWARD, and F. V. GRAU. 1953. How to keep turfgrass healthy. Pages 285-291 in: Plant Diseases, The Yearbook of Agriculture. U.S. Dept, of Agr., Washington, D.C. 939p.

49. LUKENS, R, J. 1960. Conidial production from filter paper cul­ tures of Helminthosporium vagans and Alternaria solani. Phytopathology. 50:867-868.

50. LUKENS, R. J. 1968. Low light intensity promotes melting-out of bluegrass. (Abstr.) Phytopathology. 58:1058. 15

51. LUKENS, R. J. 1970. Melting-out of Kentucky bluegrass, a low sugar disease. Phytopathology. 60:1276-1278.

52. McCALLAN, S. E. A. 1957. Effectiveness of various formulations of Bordeaux mixture in controlling early and la.te blights of tomato in greenhouse tests. Contrib, Boyce Thompson Inst. 19:157-167.

53. MEREDITH, D. 1963. Violent spore release in some Fungi Imper- fecti. Ann, Bot, 27:39-47.

54. MEREDITH, D. 1963. Further observations on the zonate eyespot fungus, Drechslera gigantea in Jamaica. Trans. Brit. Mycol. Soc. 46:201-207.

55. MEREDITH, D. 1965. Violent spore release in Helminthosporium turcicum . Phytopathology. 55:1099-1102.

56. MEREDITH, D. 1966. Airborne conidia of Helminthosporium turcicum in Nebraska. Phytopathology. 56:949-952.

57. MONTEITH, J. 1924. The leaf-spot disease of bluegrass. Bull. Green Sect. U.S. Golf Assoc. 4:172-173.

58. MORRISON, R. H. 1980. Leaf spot of Kentucky bluegrass in Minne­ sota: Misidentification of Drechslera poae as Drechslera dictyoides. In: P. 0. Larsen and B. Joyner, eds. Advances in Turfgrass Pathology. Harvest Publications, Cleveland, Ohio (in press).

59. MOWER. R. G. 1961. Histological studies of suscept-pathogen relations of Helminthosporium sativum, Helminthosporium vagans, and Curvularia lunata on leaves of Merion and common Kentucky bluegrass (Poa pratensis L.) Ph.D. Thesis, Cornell Univ., NY. 150p.

60. NEELY, D. 1970. Persistance of foliar protective fungicides. Phytopathology. 60:1583-1586.

61. NEELY, D . , and E. B. HIMELICK. 1966. Simultaneous determination of fungistatic and fungicidal properties of chemicals. Phyto­ pathology. 56:203-209.

62. TORGENSON, D. C. 1969. Determination and measurement of fungi- toxlcity. Pages 93-123 in: D. C. Torgenson, ed. Fungicides, an advanced treatise. Academic Press, London and New York. Vol. I. p.697.

63. VARGAS, J. M . , and R. D. WILC0XS0N. 1969. Some effects of temperature and radiation on sporulation by Helminthosporium dictyoides on agar media. Phytopathology. 59:1706-1712. 16

64. WAGGONER* P. E. 1973. The removal of Helminthaporium maydls spores by wind. Phytopathology. 63:1252-1255.

65. WALLIN, J. R. 1973. Spore sampling of Helminthosporium maydis race T in 1971. Plant Dis. Reptr. 57:298-299.

66. WEIHING, J. L., S. G. JENSON, and R. I. HAMILTON. 1957. Helminthosporium sativum* a destructive pathogen of bluegrass. Phytopathology. 47:744-746. Chapter I

MELTING-OUT OF KENTUCKY BLUEGRASS: SOURCE OF INOCULUM RELEASE, AND DISPERSAL OF DRECHSLERA POAE CONIDIA ABSTRACT

Populations of Drechslera poae conidia in the thatch and leaf

litter of Kentucky bluegrass turf were monitored in 1979 and 1980.

Results indicate that leaf litter and not thatch is the primary source

of inoculum in the spring. Peak populations in the leaf litter coin­

cided with periods of heavy D. poae conidium release as well as sub­

stantial increases in diseases incidence on Kentucky bluegrass in May

and June of 1980. During this period, thatch temperatures ranged

from 6 to 20 C with peak conidium production occurring at 9 to 18 C.

Moisture in the form of rainfall or heavy 8-12 hour dew periods were

plentiful during this period. Low temperatures and sporadic rainfall

limited conidium populations in April or early May, Sporulation in leaf

litter ceased in early July, 1980 as the mean daily thatch temperature

exceeded 20 C.

The diurnal release of D. poae conidia peaked between 1200 and

1400 hours. The incidence of airborne conidia began to increase at

0800 and substantial numbers of conidia were collected through 2000

hours. Few conidia were trapped in the late evening or early morning.

Discharge of conidia was associated primarily with reductions in the

moisture levels in turf microenvironment while rainfall and wind were

of minor importance. Negligible releases of conidia were noted during

periods of prolonged leaf wetness or high humidity.

19 20

The incidence of the leaf spot and crown rot phases of melting- out peaked in May and June when inoculum was plentiful. During the summer in 1979, disease severity remained high.

In September, a rapid decrease in disease Incidence was noted when conditions for rapid plant growth were favorable. The low incidence of disease was reflected in the low population of D. poae conidia in leaf litter and generally low level of D.poae conidium discharge.

INTRODUCTION

Melting out which is caused by Drechslera poae (Baudys) Shoem.

(^Helminthosporium poae Baudys, Helminthosporium vagans, Drechs., and Drechslera vagans (Drechs.) Shoem.) is an important disease of Ken­ tucky bluegrass throughout the United States (6, 7, 8, 9, 10, 11, 13,

14, 33, 34). Two distinct phases of melting-out generally develop during the growing season. The leaf spot phase, which is most visible develops in the cool, humid spring and fall, while the crown rot phase is most prominent during the summer (6, 7). Although the appearance of the crown rot phase was often sporadic In Minnesota, Morrison (33) noted that the crown rot phase was most severe in the late spring rather than during the summer.

Drechslera poae overwinters on infected plant material or leaf litter as vegetative hyphae or conidia (6, 7, 14, 16) . The primary source of inoculum, i. e., D. poae conidia, throughout the year may be thatch, leaf litter or lesions on diseased Kentucky bluegrass plants (3, 7).

Colbaugh and Beard (4) have found that sporulation of Helminthosporium spp. is poor on intact plants but may be quite heavy on leaf litter. 21

Maintenance practices such as irrigation, mowing height and rate of

fertilization may have a signficant Influence on the population of

Helminthosporium spp. conidia in bermudagrass leaf litter (3,4).

The availability of moisture in Kentucky bluegrass leaf litter also

has a profound effect on the saprophytic and parasitic activity of

Bipolaris sorokiniana ("Helminthosporium sorokinianum) (5). Sporu­

lation of B. sorokiniana was 20 to 30X higher on Kentucky bluegrass

leaf litter in drought stressed areas than in areas receiving adequate

moisture (5).

Little information concerning the release and dispersal of air­

borne conidia of any turfgrass pathogen was available. Couch (6, 7)

states that D. poae conidia were splashed onto the foliage by rain but

no data exists to back this assertion. Meredith (30) observed the diurnal periodicity of Drechslera gigantea ("Helminthosporium giganteum

over bermudagrass turf. Peak concentration of airborne I). gigantea

conidia occurred at approximately 1000 hours (30). The incidence of

airborne conidia of Bipolaris maydis ("Helminthosporium maydis) and

Exserohilum turcica ("Helminthosporium turcicium) also occurred be- ween 0900 and 1100 hours (1, 25). Meredith (30) also observed that

the incidence of airborne I), gigantea was highest on dry days following

rainfall.

Periods when the incidence of airborne Helminthosporium conidia was highest coincide with substantial changes in relative humidity,

temperature or wind velocity (1, 25, 30). Research has shown that

changes in relative humidity, infrared radiation, and wind velocities

of 3-6 m/sec Influence the removal of conidia from conldlophores 22

(1, 2, 17, 21, 23, 25, 30, 36).

The objective of this research was to evaluate the influence of weather variables on the development of the leaf spot and crown rot phase of melting-out of Kentucky bluegrass caused by Drechslera poae.

Emphasis was placed on determining the site of conidium production and the incidence of airborne D. poae conidia within a field of Ken­ tucky bluegrass throughout the growing season.

MATERIALS AND METHODS

Conidium trapping: Airborne conidia of Drechslera poae were col­ lected with a Kramer-Collins intermittant band spore sampler (G-R

Electric Manufacturing Co., Manhattan, KS) or a Burkard 7 day continuous spore sampler (Burkard Scientific Ltd., Rickmansworth, Hertfordshire,

England).

The Kramer-Colllins spore trap was adjusted to operate approxi­ mately 23 min/hr at a flow rate of 35 1/hr through an orifice 15 cm above the ground. Particulate matter was deposited on a glass slide treated with WD-40 (WD-40 Company, San Diego, CA) penetrating oil.

Samples were collected with the Kramer-Collins trap from April 10 to

July 10 plus August 27 through November 12 in 1979 and from March 17 to July 10, 1980. Slides were changed dally, stained with 0.5% cotton blue in lactophenol and examined for the presence of poae conidia.

The Burkard spore trap operated at a flow rate of 10-11 liters of ■i air/min with the orifice at 30 cm above the turf. Conidia were col­ lected on petroleum jelly coated plastic trap tapes. Weekly samples were collected with the Burkard trap from July 27 to November 2, 1979 and April 1 to July 14, 1980. Tapes collected from the trap were cut into 48 mm strips, mounted on glass slides, stained with 0.5% 23 cotton blue in lactophenol, and examined microscopically at 100X mag­ nification. 2 Both traps were installed in or around a 1.5 m enclosure in an

11 acre common Kentucky bluegrass field naturally infested with Drech­ slera poae at the ChemLawn Research Facility in Milford Center, Ohio.

The spore traps were situated around the enclosure to take advantage of the prevailing west-southwest winds. The bluegrass was maintained at a height of 3.8 to 5.1 cm, irrigated when necessary, and treated with broadleaf herbicides. The 11 acres of Kentucky bluegrass were not maintained at uniform fertility levels.

Meterological equipment: A hygrothermograph (Weather Measure

Corp., Sacramento, CA) anemometer (Weather Measure Corp.), leaf wet­ ness meter (28), pyranograph (Weather Measure Corp.) and recording rain gauge (Meterology Research, Inc., Altadena, CA) were installed in the enclosure containing the spore traps. Thatch, ambient air, and soil thatch temperature were monitored with t-type thermocouples, which were placed in Kentucky bluegrass turf 50 m from the enclosure and wired to a multipoint recorder (Leeds and Northrup Corp., North

Wales, PA). All equipment except the pyranograph and leaf wetness meter were installed on April 1, 1979 and removed on November 2, 1979.

The pyranograph and leaf wetness meter were installed in September and October, 1979, respectively. In 1980, all instruments were op­ erational on March 11, 1980, except the leaf wettness meter which was installed on April 19, 1980. Collection of weather data was termina­ ted on July 14, 1980.

Inoculum source: Two thatch samples were collected bi-weekly from March 26 to November 21, 1979 and March 17 to July 10, 1980 with 24 a 2.5 cm soil probe from a 95 tn^ plot divided into 64 subplots.

Leaf litter samples were collected using a clawed hand tool along with the thatch samples from 4 sub plots from August 16 to November

21, 1979 and from March 15 to July 10, 1980. Before processing, each sample was air dried under a bank of fluorescent lights.

The number of Drechslera poae and Bipolaris sorokiniana conidia in the thatch and leaf litter samples determined using the mineral oil flotation technique developed by Ledingham and Chinn (27) and further modified by Colbaugh (25). Ten grams (dry weight) of thatch or five grams (dry weight) of leaf litter were placed in a flask con­ taining 100 ml distilled water. The flask was shaken vigorously for

3 minutes and the soil-water suspension was filtered through a

200 mesh screen to remove debris. The filtrate was poured into a graduated cylinder, brought to a volume of 100 mis distributed into

25 ml aliquots and centrifuged at 480 xg. The supernatant was dis­ carded and the pellet was mixed with 5 ml of light mineral oil. Ap­ proximately 20 ml of double distilled water was emulsified with the mineral oll-soil suspension with a vortex mixer. The emulsion was poured into a glass petri dish and allowed to settle. A total of four plates were prepared from each 10 g thatch or 5 g leaf litter sample.

Six fields at 60X magnification of each plate were examined with a diseccting microscope and the number of D. poae conidia was recorded.

The area of the microscope field and petri plate were calculated, and the total number of conidia in the four plates totaled. The population of D, poae conidia/g dry weight of sample was calculated by dividing the total number qf conidia extracted by the weight of the sample (5 25 or 10 g). Drechslera poae conidia, as well as, those of several

Bipolaris spp. were found at the oil-water interface rather than floating in the mineral oil as noted by Ledingham and Chinn (27).

Disease incidence: The incidence of the crown rot and leaf spot phase of melting out was monitored weekly from March through November in 1979 and March through July, 1980. Disease incidence was expressed as a percentage of the 25 plants inspected displaying disease symptoms.

RESULTS

Populations of D_. poae in the thatch of Kentucky bluegrass turf were quite low and never approached the populations in the leaf litter

(Figures 1.1-1.3). Never more than 35 conidia/g dry weight thatch were detected in any thatch sample collected in 1979 or 1980 (Figures 1.1-

1.3). Although variable, populations of conidia in thatch were higher in the spring of 1979 than in the summer or fall (Figures 1.1, 1.2).

In 1980, populations did not exceed 20 conidia/g dry weight thatch throughout the spring and dropped to zero by July (Figure 1.3).

Inoculum source: Leaf litter consisting of decomposing Kentucky bluegrass leaf fragments was apparently the primary source of Drechslera poae conidia in the field (Figures 1.2, 1.3). In 1979, no conidia were detected in leaf litter collected from August through most of October

(Figure 1.2). A sharp increase in the population of Drechslera poae conidia was noted in a leaf litter sample collected in late October but the population of conidia immediately dropped to near zero in early

November (Figure 1.2). High populations of D. poae in 1980 were noted in leaf litter collected from mid-May through late June (Figure 1.3).

Conidium populations in leaf litter from March to May were highly 26

Figure 1. 1 Populations of Drechslera poae conidia in Kentucky bluegrass thatch from March 26 to July 16, 1979. Conidia/g dry weight substrate 400. . 500. _ 200 300. - 600. . 100 700. 800. . ..

/6 / 41 41 42 51 5/8 5/1 4/24 4/19 4/10 4/5 3/26 » ^ hth—— — Thatch ---- # /4 /2 /9 / 61 62 72 /1 7/16 7/11 7/2 6/27 6/18 6/5 5/29 5/22 5/14 1979 ___1

--i ------^ - ro *^1 Figure 1.2 Populations of Drechslera poae conidia in Kentucky bluegrass thatch and leaf litter from July 25 to November 2, 1979. Conidia/g dry weight substrate 400. 600. 200 300. 500. 800. 0 0 1 700. . . /5 /1 / 81 82 82 9/4 8/28 8/21 8/14 8/7 7/31 7/25 1 ------hth - litter Leaf — Thatch 1 ----- 1 -----

------9/10 1979 /7 /4 01 08 01 1/2 11/2 10/22 10/15 10/8 10/1 9/24 9/17 30 variable (Figure 1.3) but lower than populations in May and June

(Figure 1.3).

Thatch temperatures apparently had an important impact on the population of D. poae conidia in Kentucky bluegrass leaf litter during the spring 1980. Populations of 400 to 700 D. poae conidia/g dry weight leaf litter were observed at mean thatch temperatures between 9 to 18 C while populations between 0 to 300 conidia/g dry weight leaf litter were noted at 5 to 9 C (Figure 1.3, 1.4). Populations of coni­ dia in leaf litter dropped to nearly zero as the mean temperature exceeded 20 C in July (Figures 1.3, 1.4). Moisture in the form of rain and dew was plentiful during May and June (Figure 1.4). Sub­ stantial increases in D. poae populations in leaf litter were even noted during the dry period from mid April through early May. Quite possibly, heavy dews during this period provided adequate moisture for the initiation of D. poae conidium production.

Note: Information presented in Figure 1.5 to 1.9 concerning conidium formation and discharge is representative of the data col­ lected in 1979 and 1980.

Factors influencing sporulation of Drechslera poae in the field:

It was difficult to pinpoint the minimal period of leaf wetness required for sporulation in this study. The period of leaf wetness of Kentucky bluegrass leaves, sheaths, or leaf litter varied up to several hours per day. Sporulation often followed periods of rainfall or irrigation

(Figure 1.6-1.9). Moderate releases of D. poae conidia on May 13 and

21, 1980, were preceded by rainfall which kept plants and leaf litter wet at least 36 hours prior to conidium release (Figure 1.6, 1.7). Figure 1.3 Populations of Drechslera poae conidia in Kentucky bluegrass thatch and leaf litter from March 17 to July 10, 1980. Conidia/g dry weight substrate 600, 400 500. 200 300 800 100 700. Leaf litter Leaf Thatch 1980 1 10 w N> Figure 1.4 Summary of weekly total rainfall and mean thatch temperature from April 4 to July 10, 1980 at the Chemlawn Research Facility, Milford Center, Ohio. Mean thatch temp. o 20 25. 10 15. 5. _ . . / 41 41 42 52 / 51 52 53 66 /3 /8 /5 / 7/10 7/3 6/25 6/18 6/13 6/6 5/30 5/23 5/16 5/9 5/2 4/25 4/18 4/11 4/4 Rainfall - Rainfall ~ Temperature X

1980

Weekly rainfall 35

Approximately 1.3 cm of irrigation water was applied to the Kentucky bluegrass ranges at the ChemLawn Research Center on the afternoons of

May 15 and June 13, 1979 (Figures 1.8, 1.9). Nearly 48 hours later, on May 17 and June 15, moderate releases of D. poae conidia were noted

(Figure 1.8, 1.9).

In the fall of 1979, conditions were generally favorable for sporulation yet populations of conidia in the thatch were low and few airborne conidia were collected with the spore traps (Figures 1.2, 1.3).

Excessive moisture in the leaf litter may have either suppressed sporulation of D. poae on leaf litter or prevented release of D. poae conidia.

Factors Influencing the release of D. poae conidia: Release of

D. poae was influenced by drying of leaf- surfaces, wind and possibly

irrigation. Several examples of each release pattern will be discussed.

Depletion of moisture In the turf microclimate which can be Il­

lustrated by sharp decreases in the relative humidity or drying of

plant surfaces was associated with I). poae conidium release (Figures

1.5, 1.6). Discharge of D. poae conidia generally followed the dry­

ing of sheath surfaces or substantial drops in relative humidity

(Figures 1.5, 1.6, 1.8). Few airborne conidia were collected on days when the duration of leaf wetness exceeded 20 hours per day (Figures

1.6, 1.7).

Conidium release often occurred during the time of day when the wind velocity was approximately 0.5 to 2.0 m/sec 15:cm above the turf

(Figures 1.5-1.7). On May 13, 1980, at 2000 hours, the peak incidence of airborne I). poae conidia occurred when the wind velocity reached

5.8 m/sec (Figure 1.7). Figure 1.5 Influence of weather variables on the production and release of Drechslera poae conidia within a Kentucky bluegrass field from June 11-13, 1980 (LW=leaf wetness). E 3 0

U> 2 4 6 8 10 12 1416)6 202224 2 4 6 6 10 (2 14 16 1 8 2 0 2 2 2 4 24 6 8 (0 12 )4 16 (8202224 --j (hours) June 11 June 12 June 13 1980 Figure 1.6 Influence of weather variables on the production and release of Drechslera poae conidia within a Kentucky bluegrass field from May 20-22, 1980 (LW-leaf wetness, WS=wind velocity). Conidia/m 3 Temp C° 20 30 50 40 20 May20 a 21May (hours) T7V. 7///////77y73 7 y 7 7 / / / / / / / 7 __ May22 X 0 8 9 1 20 40 0 6 ' 0 8 100 Figure 1.7 Influence of weather variables on the production and release of Drechslera poae conidia within a Kentucky bluegrass field from May 12-14, 1980 (LW-leaf wetness). 25 20 15

10

5

showers— I 50 40

30 20

10

2 4 6 8 10 12 14 16 18 202224 2 4 6 8 10 12 14 16 18 2 0 2224 2 4 6 8 (hours) May 12 May 13 May 14 1980 42

D. poae conldia release Increased substantially following periods

of Irrigation. Substantial number of D. poae conldia were collected when 1.3 cm of water was applied to the Kentucky bluegrass turf at

1100 hours on May 16, 1979 (Figure 1.9). Simultaneously, a large

decrease in relative humidity was also noted (Figure 1.9). The

release of D. poae conidia did not increase following rainshowers

in 1979 or 1980.

Diurnal release of D. poae conidia; The incidence of airborne

D. poae conidia was much higher during daylight hours (0800 to 2000

hours) than at night (2000 to 0800 hours) (Figures 1.10, 1.11).

Collection of D. poae conidia which often began between 0800 and 1000 was associated with rapid changes in moisture levels in the turf microclimate in the morning, wind velocities in excess of 2 m/sec, or

in several cases irrigation (Figures 1.5-1.11). In 1979 and 1980, peak

concentration of I), poa airborne conidia occurred between 1200 to

1400 hours (Figures 1.10, 1.11). Substantial numbers of conldia were

trapped between 0800 and 1800 hours (Figures 1.10, 1.11). Discharge

5.* P°ae conidia was low between 2000 and 0800 hours, when leaf sur­

faces and leaf litter were wet and wind velocities quite low (Figures

1.5-1.11).

Seasonal distribution of airborne Drechslera poae conidia: The

incidence of airborne D. poae conidia was highest in the spring when

ambient air and thatch temperatures were generally mild and moisture was readily available (Figures 1.4, 1.12, 1.14). In 1979, peak in­

cidence of 30-80 airborne D. poae conidia nrVday was noted between

May 14 and 28 and June 4 to 24 (Figure 1.12). In late April and early Figure 1.8 Influence of weather variables on the production and release of Drechslera poae conidia within a Kentucky bluegrass field from June 13-15, 1979. 100

\ I 30 Z' 80 0 x V------V ------A v 60 St 1 2 0 ______/ x i * 10 — 40 ! 20 7 7 7 5 30 irrigation^ * 25 CO E 20 * 5 15 o 10

1 1 1 1 1 1 1 1 V T -M J. 1 1 1 i 1 1 2 4 6 8 10 12 14 16 18 20 2224 2 4 6 8 10 12 14 16 18 20 2224 2 4 6 8 10 12 14 16 18 2 0 2 2 2 4 (hours) June 13 June 14 June 15 1979 45

Figure 1.9 Influence of weather variables on the production and release of Dreehslera poae conidia within a Kentucky bluegrass field from May 15-17, 1979. too

- / 80

20 60 %RH 40 20

30 Showers v irrigation 25 E 20

2 4 6 8 10 12 W 16 18 202224 2 4 6 6 10 12 14 16 18 20 2224 2 4 6 8 1012 14 16 18 20 2224 (hours) ST' May 15 May 16 May 17 1979 Figure 1. 10 Diurnal periodicity of airborne Drechslera poae conidia collected within a Kentucky bluegrass field with a Kramer-Collins spore trap from April 17 to November 2, 1979. 0200 0400 0600 0800 1000 1200 1400 1600 1800 2000 2200 2400 Hours 1979 (April 17-November 2) .c- CO Figure 1.11 Diurnal periodicity of airborne Drechslera poae conidia collected within a field of Kentucky bluegrass with a Burkard spore trap from April 1 to July 6, 1980. 0200 0400 0600 1000 1200 1400 1600 1800 2000 2200 2400 Hours 1980 (April 1-July 6)

U l o 51

May, the total number of conldia collected was lower than levels noted in June (Figure 1.12). The incidence of airborne conidia remained low from

July to November ranging from 8-40 airborne conidia m^/day (Figure 1.13).

In 1980, the spore traps were fully operational several weeks earlier than in the spring of 1979 (Figures 1.12, 1.14). With the exception of the period between April 7 to 13, the incidence of air­ borne D. poae conidia was quite low in April and early May (Figure 1.14).

Number of airborne conidia increased between May 12 to 18 to 100-120

I), poae conidia/tn^/wk (Figures 1.4, 1.14). Peak concentration of

150-300 airborne D. poae conidia/m^/wk were noted between June 2 and

29 with moderate concentrations being recorded into early July (Figure

1.14). The incidence of airborne D. poae dropped sharply in mid-July to nearly zero as temperatures in the turf microenvironment regularly exceeded 20 C (Figures 1.4, 1.14). Collection of D. poae conidia with the Burkard spore trap was interrupted between June 19 and June 25.

Disease incidence: In 1980, a close relationship between disease incidence and periods of conidium release was observed. Lesion develop­ ment was observed 5 to 14 days after a substantial conidium release

(Figures 1.14, 1.16). For example, on April 7 a cloud of D. poae conidia with a density of 8 to 33 conidia/mJ was detected for several hours.

Approximately 14 days later, substantial increases in the incidence of the leaf spot and crown rot phase of melting out were recorded. Lesion formation was faster during May and June as temperatures in the turf microclimate increased (Figure 1.16). In mid-May, substantial in­ creases in disease incidence occurred within 5-7 days of moderate coni­ dium releases (Figures 1.14, 1.16), Figure 1. 12 Seasonal distribution of airborne Drechslera poae conidia collected within a field of Kentucky bluegrass with a Kramer-Co11ins spore trap from April 22 to July 29, 1979 (ND=no data). Conidia/m 60. 30. 50. 80. 10 20 70. . . /9 / 51 52 52 63 /0 /7 /4 / 78 /5 7/29 7/15 7/8 7/1 6/24 6/17 6/10 6/3 5/28 5/20 5/13 5/6 5/7 4/29 4/30 4/22 /4 /1 /9 / 61 61 62 7/2 6/25 6/18 6/11 6/4 5/29 5/21 5/14 99 w 1979 ND 7/167/9 ND Figure 1.13 Seasonal distribution of airborne Drechslera poae conidia collected within a field of Kentucky bluegrass with a Kramer-Collins and Burkard spore traps from July 30 to November 2, 1979 (ND=no data). Conidia/m3 AO. 60. 30. 80. 50. 70. 0 2 10 .- . 8/5 7/30 _ ND l _ 8/12 8/6 rmrClissoe trap spore Kramer-Collins ND ukr pr rp ■ trap spore Burkard I _ / 9/9 9/2

/0 9/178/279/10 9/3 /6 9/23 9/16 1979 /1 07 01 1/1 02 11/2 10/27 10/21 10/28 10/14 10/7 10/22 10/15 9/31 10/8 10/1 9/24 Ui Figure 1.14 Seasonal distribution of airborne Drechslera poae conidia collected within a field of Kentucky blue- grass with a Burkard spore trap from March 31 to July 14, 1980 (Note: collection of conidia was interrupted between June 19 to June 25). A data interrupted + ! \ / \

v+

5/19 5/26 6/2 6/9 6/16 6/23 6/30 7/7

5/25 6/1 6/8 6/15 6/22 6/29 7/6 7/14 1980 in ■^1 Figure 1.15 Incidence of leaf spot and crown rot phases of melting-out of Kentucky bluegrass caused by Drechslera poae in 1979. Disease incidence was expressed as the percentage of 25 plants inspected weekly for the presence of the leaf spot or crown rot phases of melting-out. 100. Leaf spot ■*— 90. / Crown rot — - 80. J

70. 41 O c / 60, u G 50. 4) (0 <0 $ 40

30.

2 0 .

10.

^ __X> X X 4/1 4/15 5/1 5/15 6/1 6/15 7/1 7/15 8/1 8/15 9/1 9/15 10/1 10/15 11/1 11/15 1979

vO Figure 1.16 Incidence of the leaf spot and crown rot phases of melting-out of Kentucky bluegrass caused by Drechslera poae from March 24 to July 10, 1980, Disease incidence was expressed as the percentage of 25 plants inspected weekly for the presence of the leaf spot or crown rot phases of melting-out. Disease incidence 100 40. 10 20 80. 90. 30. 50. 60. 70. . . . /4 / 41 42 42 51 51 52 53 66 /3 /8 /5 / 7/10 7/3 6/25 6/18 6/13 6/6 5/30 5/23 5/16 5/12 4/28 4/23 4/11 4/1 3/24 ef spot Leaf Crown rot Crown

1980 O' 62

A poor relationship was noted between disease incidence and the seasonal incidence of airborne D_. poae conidia in the spring of 1979

(Figures 1.12, 1.15). Substantial increases in disease incidence were observed in April despite the absence of airborne j). poae conidia.

Disease incidence did continue to increase when airborne conidia were plentiful in May and June (Figures 1.12, 1.15). Only small decreases in the incidence of the leaf spot and crown rot phases of melting out occurred during July and August when the plants were generally dormant.

Sharp decreases in disease incidence were noted in September through

November when inoculum levels were low (Figures 1.13, 1.15).

It is important to note that the development of the leaf spot and crown rot phase was simultaneous through the spring of 1979 and

1980 (Figures 1.15, 1.16). The crown rot phase was quite prominent and often serious during the spring and early summer in 1979 and 1980 but severe melting-out was never observed. Evidently, environmental conditions favorable for the development of severe melting-out of

Kentucky bluegrass were never present during 1979 or 1980. 4

DISCUSSION

Kentucky bluegrass leaf litter was the most important source of inoculum in the spring, while thatch and lesions on diseased leaves were minor sources. High populations of conidia in leaf litter coin­ cided with peak concentrations of airborne D. poae conidia and preceded substantial increases in the incidence of the leaf spot and crown rot phases of melting out (Figures 1.13, 1.15, 1.16). Colbaugh and Beard

(A) and Colbaugh and Endo (5) have noted that conidium production of

Helminthosporium spp on bermudagrass leaf litter and B. sorokiniana on 63

Kentucky bluegrass leaf litter was also quite heavy. Populations of

D. poae which peaked between 600-700 conidia/g dry weight leaf litter did not approach the population levels of Helminthosporium spp. noted by Colbaugh and Beard (4) in leaf litter from tall fescue, bermudagrass, and St. Augustinegrass turf in Texas.

Couch (7) noted that thatch, i.e., crowns, roots and rhizomes of

Kentucky bluegrass plants, in addition to leaf litter served as a site of inoculum production. In this study even during periods of peak conidium populations in Kentucky bluegrass leaf litter, populations of

poae conidia in thatch were quite low (Figure 2.3). Conidia of D. poae found in the thatch probably were deposited there after being dislodged from leaf litter.

Intact diseased plants were not evaluated as a source of Inoculum in the field. Colbaugh and Beard (4) observed that sporulation of

Helminthosporium spp. rarely occurs on plants in the field. Halisky and Funk (14) observed poor I). poae sporulation on lesions on Kentucky bluegeass leaves collected between May and July, periods of peak I), poae airborne conidium incidence (Figures 1.12, 1.14). Laboratory studies have indicated that sporulation of D. poae is 4 to 10 times higher on excised diseased leaves than on diseased leaves collected from intact plants (Table 2.2).

It is difficult to pinpoint in this study the duration of free moisture required for D. poae conidium formation on infested plant material in the field. Heavy releases of D. poae conldia were observed following periods of rainfall or heavy dews when the leaf litter re­ mained continuously moist for at least 36 hours (Figures 1.6, 1.7).

Laboratory results confirm field observations which indicate that 64

24 to 36 hours of continuous free moisture are required for conidiophore development and conidiura formation on excised Kentucky bluegrass leaves (Chapter II).

Conidium formation was not limited to periods of continuous free moisture in excess of 24 hours. Sporulation of D. poae also occurred during dews of 8 to 10 hours duration on several successive nights

(Figure 1.5). The number of dew periods on successive evenings re­ quired for conidium production is unknown. Leach et al (26) noted that the presence of free moisture, i.e., dew or rainfall, for 10 to 12 hours on corn leaves at night favored Exserohilum turcica conidium formation. However, E_. turcica often produces successive crops of conidia for several days on the same conidiophores thereby reducing the time required for the production of conidia on corn leaves. Pro­ duction of successive crops of D. poae conidia on mature conidiophores may not occur. Competition between D. poae and other saprophytes on leaf litter probably limits conidium production.

The level of D. poae sporulation in the field whcih reflected by the conidiura population in leaf litter was heaviest at mean thatch temperatures of 9 to 18 C (Figures 1.3, 1.4). Below 9 C, sporulation on leaf litter was light. As the daily mean thatch temperatures ex­ ceeded 20 C, sporulation of I), poae rapidly decreased in leaf litter.

Field and growth chamber data concerning influence of temperature

£.. poae sporulation were in agreement. Sporulation of IK poae on excised Kentucky bluegrass leaves peaked at 12 and 18 C but was light at 6 C (Figure 2.10). Vargas and Wilcoxson (35) also noted heavy D. poae sporulation at 15 and 18 C in vitro. Sporulation of

D, poae began to decrease at 24 C on excised leaves (Figure 2.10). 65

However, poor sporulation was noted by Vargas and Wilcoxson (35) at

21 C.

Melting-out of Kentucky bluegrass has been recognized as a

major disease throughout the growing season in the northern United

States (6, 7, 14). Usually, the leaf spot phase developed in the

spring and fall while the crown rot phase predominates during the

summer (6, 7). In the spring of 1979 and 1980 simultaneous increases

in the incidence of the leaf spot and crown rot phases of melting-out

were noted (Figures 1.15, 1.16), Disease development followed in­

creases in the population of D. poae conidia in leaf litter and in­

cidence of airborne D_. poae conidia. Little increase in disease in­

cidence was observed from late May to early June when inoculum was

plentiful because leaf and crown lesions were found on nearly all Ken­

tucky bluegrass plants examined.

Severe melting-out of Kentucky bluegrass did not develop during

the summer of 1979 or 1980. Although close inspection of Kentucky

bluegrass plants in July and August did reveal that some were severely

diseased, overall turf quality was good. Decreases in disease inci­

dence were small in 1979 but quite large in July 1980 (Figures 1.15,

1.16). Morrison (33) noted that the occurrence of melting-out, though

sporadic, was most severe In the late spring rather than during the

summer.

In the fall of 1979, environmental conditions were ideal for dis­

ease development yet sporulation of D. poae and disease incidence were low. Other authors have noted the development of the leaf spot

phase in the fall (6, 7, 14). It is quite possible that heavy rainfall

throughout the fall suppressed conidium formation. Langenberg et al (20) have observed that prolonged periods of rain or wet leaves, cool temperatures, and high wind velocities inhibited sporulation of Alter- naria dauci on carrot. Sporulation of B^ sorokiniana was substantially higher on Kentucky bluegrass leaf litter from drought stressed turf than from turf receiving adequate moisture (5). Evidently alternating wet-dry cycles favored heavy sporulation of sorokiniana (5), and it is quite possible thati the same pattern of wet-dry cycles influenced sporulation of D. poae on leaf litter.

The discharge of D_. poae conidia follows a diurnal pattern with peak concentrations of airborne conidia occurring between 1200 and

1400 hours. This peak is several hours later than those reported for

D. gigantea and E. turcica (25, 30, 32). The incidence of airborne conidia begins to rise at 0800 and discharge of conidia often continued until 2000 hours (Figures 1.10, 1.11). Few conidia were collected at night or on days when wind velocity was low and moisture levels high.

Release of D. poae conidia was associated with substantial drops in the level of moisture, i.e., relative humidity or leaf wetness. Dis­ charge of conidia followed sharp drops in the ambient relative humidity by 1 to 2 hours on a number of occasions (Figures 1.5-1.9). A close relationship between conidium release and evaporation of surface mois­ ture from plant surfaces and leaf litter was noted in 1980. Discharge of D. poae conidia may have been delayed until late afternoon on days following rain showers because of excessive moisture levels in leaf litter. Meredith (32) and Leach et al (25) have observed the Importance of changes in the relative humidity and evaporation of free moisture on leaf surfaces as causes of conidium release. Release of I), poae 67 conidia was negligible during prolonged periods of high relative humidity or leaf wetness which occurred on rainy or cloudy days, as well as, at night. Prolonged periods of high relative humidity or leaf wetness had a similar influence on the release of E. turcica and

2.* glgantea (25, 32).

Two theories have been developed to explain the moisture related violent discharge of _E. turcica and D. gigantea conidia (22, 24, 30,

31). Meredith (30, 31) attributed the forcible discharge of E. turcica, and D. gigantea conidia to mechanical stress as well as production of gases within the conidium and conidiophore after desiccation. Leach

(22, 24) demonstrated that forcible discharge of E. turcica conidia is caused by the repulsion of unipolarly charged conidia from a charged surface of the same polarity.

It is difficult to evalute the role of wind on I), poae conidium discharge. Although light winds with velocities of 0.5 to 2.0 m/sec are sufficient to keep large conidia such as D. poae airborne, it is questionable whether light winds could dislodge conidia from conidio­ phores (12).

Some research has centered on determining the wind velocity re­ quired to removed Blpolaris maydis conidia from c o m leaves (1, 2, 36).

Waggoner (36) noted that wind velocities of 3 to 7 m/sec were required for substantial release of B^. maydis conidia from c o m leaves in a wind tunnel. Further work in the laboratory by Aylor and Parlange (2) has shown that winds of less than 2 m/sec/hr removed significant num­ bers of B_. maydis conidia from c o m leaves. During very brief periods of time, the velocity of wind puffs exceeded 4.5 m/sec over the coni­ dium bearing surface although the mean wind velocity did not exceed 68

2 m/sec/hr (2). Aylor and Lukens (1) observed substantial releases of IJ. maydis conidia from corn leaves in the field at 0.5 to 1.4 m/sec.

Since _B. maydis conidia were collected between 0900 and 1100 hours, conidium release may have been caused by abrupt changes in ambient relative humidity which generally occurs at this time of day (1).

The question of the influence of wind velocity on IK poae conidium discharge must be answered using wind tunnel studies rather than relying solely on field data.

Discharge of D. poae conidia was not observed during light showers or heavy thunderstorms although light irrigation may cause conidium release. Rapid washout of airborne D. poae conidia during rain showers may have prevented detection of signficant discharges of conidia. Mere­ dith (32) also noted that the incidence of airborne Z. turcica conidia was quite low during periods of rainfall. However, later studies have indicated that rainfall may trigger significant releases of E. turcica conidia (25).

Little effort has been made to study the epidemiology of nelting­

out (6, 7, 13, 14). This study certainly does not provide answers to all epidemiologically-related questions but it does provide the ground work for further research. Further work should be directed at identifying the influence of chemicals such as fertilizers, herbicides, and fungicides on population dynamics of I), poae in the turf micro­ environment as well as further refining of techniques used for collecting and analyzing field data. 69

Melting-out is controlled primarily by using a preventive fungi­ cide spray program in combination with cultural control procedures in the spring and fall. Integration of field data concerning environment­ al influences on D. poae population and disease development into pre­ ventive fungicide programs will allow the turf manager to improve the timing of fungicide applications . The ultimate objective of this research program is the development of a program integrating cul­ tural and chemical control measures for effective control of melt­ ing-out of Kentucky bluegrass.

« LITERATURE CITED

1. AYLOR, D. E. and R. J. LUKENS. 1974. Liberation of Helminthos­ porium maydis spores by wind in the field. Phytopathology. 64:1136-1138.

2. AYLOR, D. E. and J. Y. PARLANGE. 1975. Ventilation required to entrain small particles from leaves. Plant Physiol. 56:97-99.

3. BEARD, J. B. 1973. Turfgrass: science and culture. Prentice- Hall, Inc., Englewood Cliffs, NJ. 658p.

4. COLBAUGH, P. F., and J. B. BEARD. 1980. Influence of cultural practices on conidial production by Helminthosporium spp. on improved bermudagrass. In P . 0. Larsen and B. G. Joyner, eds. Advances in Turfgrass Pathology, Harvest Publications, Cleve­ land, Ohio (in press).

5. COLBAUGH, P. F. and R. M. ENDO. 1974. Drought stress: an impor­ tant factor stimulating the development of Helminthosporium sativum on Kentucky bluegrass. Pages 328-333 In: E. C. Roberts (ed). Proc. 2nd Internatl. Turfgrass Res. Conf, American Soc. Agronomy.

6 . COUCH, H. B. 1957. Melting-out of Kentucky bluegrass, its cause and control. Golf Course Reptr. 25:5-7.

7. COUCH, H. B. 1973. Diseases of turfgrasses. Krieger Publishing Co., Huntington, NY. 348p.

8 . DRECHSLER, C. 1922. A new leaf spot of Kentucky bluegrass caused by an undescribed species of Helminthosporium. (Abstr.). Phytopathology. 13:35.

9. DRECHSLER, C. 1930. Leaf spot and foot rot of Kentucky bluegrass caused by Helminthosporium vagans. J. Agr. Res. 40:447-456.

10. ELLIOT, E. S. 1962. The effect of soil fertilization on develop­ ment of Kentucky bluegrass diseases. (Abstr.). Phytopathology. 52:1218.

11. ELLIOT, E. S. 1962. Disease damage on forage grasses. Phyto­ pathology. 52:448-451.

12. GREGORY, P. H. 1961. The microbiology of the atmosphere. Leonard Hill, London. 251p.

70 71 13. HAGAN, A. K. and P. 0. LARSEN. 1979. Research on Helminthosporium melting-out of Kentucky bluegrass. Proc. Ohio Turfgrass Conx. 1979:46-54.

14. HAL1SKY, P. M. and C. R. PUNK. 1966. Environmental factors af­ fecting growth and sporulation of Helminthosporium vagans and its pathogenicity to Poa pratensis. Phytopathology. 56:1294-1296.

15. HIRST, J. M. 1952. An automatic volumetric spore trap. Ann. Appl. Biol. 39:257-265.

16. HORSFALL, J. G. 1930. A study of meadow-crop diseases in New York. Cornell Univ. Agr, Expt. Sta. Man. 130. 139p.

17. KENNETH, R. 1964. Conidial release in Helminthospora. Nature. 202:1025-1026.

18. KRAMER, C. L., and J. M. PADY. 1966. A new 24 hour spore sampler. Phytopathology. 56:517-520,

19. KRAMER, C. L., M. G. EVERSMEYER, and T. I. COLLINS. 1976. A new 7-day spore sampler. Phytopathology. 66:60-61.

20. LANGENBERG, W. J., J. C. SUTTON, and T. J. GILLESPIE. 1977. Relation of weather variables and periodicities of airborne spores of Alternaria dauci and other fungi. Phytopathology. 67:879-883.

21. LEACH, C. M. 1975. Influence of relative humidity and red-infra­ red radiation on violent spore release by Drechslera turcica and other fungi. Phytopathology. 65:1303-1312.

22. LEACH, C. M. 1976. An electrostatic theory explains violent spore liberation by Drechslera turcica and other fungi. 68:63-68.

23. LEACH, C. M. 1980. Influence of humidity and red-infrared radia­ tion on spore discharge by Drechslera turcica - additional evidence. Phytopathology. 70:192-196.

24. LEACH, C. M. 1980. Evidence for an electrostatic mechanism in spore discharge by Drechslera turcica. Phytopathology. 70:206-213.

25. LEACH, C. M . , R. A. FULLERTON, and K. YOUNG. 1977. Northern leaf blight of maize in New Zealand. Release and dispersal of conidia of Drechslera turcica. Phytopathology. 67:380-387.

26. LEACH, C. M . , R. A. FULLERTON, and K. YOUNG. 1977. Northern leaf blight of maize in New Zealand. Relationship of Drechslera turcica airspora to factors influencing sporulation, conidium development, and chlamydo$pore formation. Phytopathology. 67:629-636. 72

27. LEDINGHAM, R. J. and S. H. F. CHINN. 1955. A flotation method for obtaining spores of Helminthosporium sativum from soil. Can. J. Bot. 33:298-303.

28. LUKENS, R. J. 1968. Low light intensity promotes melting-out of bluegrass (Abstr.). Phytopathology. 58:1058.

29. MELCHING, J. S. 1974. A portable self-contained system for the continuous electronic recording of moisture conditions on the surfaces of living plants. ARS-NE-42, Agricultural Research Science, USDA. 13p.

30. MEREDITH, D. 1963. Further observations on the zonate eyespot fungus, Drechslera gigantea in Jamaica. But. Mycol. Soc. Trans, 46:201-207.

31. MEREDITH, D. 1965. Violent spore release in Helminthosporium turcicum. Phytopathology. 55:1099-1102.

32. MEREDITH, D. 1966. Airborne conidia of Helminthosporium turcicum in Nebraska. Phytopathology. 56:949-952.

33. MORRISON, R. H. 1980. Leaf spot of Kentucky bluegrass in Minne­ sota: Misidentification of Drechslera poae as I), dictyoides. In P. 0. Larsen and B. G. Joyner, eds. Advances in Turfgrass Pathology, Harvest Publications, Cleveland, Ohio (in press).

34. MOWER, R. G. 1961. Histological studies of suscept-pathogen relationships of Helminthosporium sativum, Helminthosporium vagans and Curvularia lunata on leaves of ’Merlon' and com­ mon Kentucky bluegrass (Poa pratensis L.). Ph.D. Thesis, Cornell Univ., NY. 150p.

35. VARGAS, J. M. and R. D. WILCOXSON. 1969. Some effects of tem­ perature and radiation on sporulation by Helminthosporium dictyoides on agar media. Phytopathology. 59:1706-1712.

36. WAGGONER, P. E. 1973. The removal of Helminthosporium maydis spores by wind. Phytopathology. 63:1252-1255. Chapter II

INFLUENCE OF TEMPERATURE ON DRECHSLERA POAE ACTIVITY AND LESION DEVELOPMENT ON KENTUCKY BLUEGRASS LEAVES ABSTRACT

Pots of Kentucky bluegrass plants were inoculated with a suspension of D. poae conidia and incubated at 6 , 12, 18, 24, and 30 C. Fungal pre-penetration events were monitored 2, 4, 6 , 8 , 13, 16, 24, 32, and

48 hours after inoculation using the collodion leaf impression technique.

Penetration and colonization of leaves was observed in cleared tissue.

Conidiura germination began 4 hours after inoculation at all temperatures.

Initially, the rates of conidium germination, germ tube elongation, and appressorium formation were often slower than the rate of development at 18 or 24 C but few differences were noted 48 hours after inoculation.

Although the number of successful penetrations 24 hours after inocula­ tion was relatively high at all temperatures, ramification of primary and secondary hyphae was poor at 6 and 30 C. Lesion expansion which was rapid at 12 to 24 C and poor at 6 and 30 C reflected the rate of hyphal' development in host tissue. Conidium production of D. poae on excised leaves is rapid and profuse at 12 to 24 C but sparse at 6 and 30 C. The level of D. poae sporulation is much higher on excised leaves than on Intact leaves from Kentucky bluegrass plants.

INTRODUCTION

Development of melting-out on Kentucky bluegrass caused by Drech­ slera poae (Baudys) Shoem. (■Helminthosporium poae Baudys, Helminthos­ porium vagans Drechs., Drechslera vagans (Drechs,)Shoem.) is greatly influenced by temperature (3, 4, 6 , 7). The leaf spot phase of melt­ ing out is usually observed in the spring and fall while the crown

74 75 rot phase is often most severe in the late spring and summer (3, 4, 9,

16, 17, 18). Field observations of leaf spot and crown rot lesion de­ velopment indicate that inoculum production and subsequent infections of bluegrass plants are favored by the cool moist conditions in the spring and fall (3, 4, 9). The activity of I), poae in plant tissue is not limited to cool temperatures. Bean and Wilcoxson (1) noted moderate reductions of 'Park* Kentucky bluegrass seedling stands ino­ culated with D. poae at temperatures up to 29 C.

There has been some confusion concerning the identity of the

Drechslera species causing leaf spot and crown rot of Kentucky bluegrass in the midwest. Bean and Wilcoxson (1) reported that D. dictyoides and not D. poae was the most important pathogen of Kentucky bluegrass in Minnesota. As a result of his failure to isolate D. dictyoides from Kentucky bluegra'ss, Morrison (17) studied the etiology of melting- out of Kentucky bluegrass. Morrison (17) concluded that IK poae had been misidentifled as I), dictyoides by Bean and Wilcoxson (1) and Vargas and Wilcoxson (21). Therefore, reference to the above mentioned re­ port (1, 21) will be assumed to relate to I), poae rather than IK dictyoides.

Conidium germination of IK poae on nutrient media or leaf tissue was quite rapid (10, 18). Within 18-24 hours of inoculation at tempera­ tures from 3 - 35 C, 85-98% of the D. poae conidia examined had germina­

ted (10, 18). Germ tube development was nearly completed within 18 hours on tips of 55% of the germ tubes on bluegrass leaves at 24 to 30 C (18).

Halisky and Funk (9) noted that penetration by I), poae of a suscepti­ ble Kentucky bluegrass cultivar 'Newport' occurred within 6 hours of in­ oculation while penetration of the resistant cultivar 'Dwarf' was delayed

12 hours after inoculation. Development of primary hyphae of I), poae 76 was noted in the substomatal cavities or epidermal cells of either

susceptible or resistant Kentucky bluegrass cultivars by Mower (18)

18 to 24 hours after inoculation. The greatly expanded globose, hya­

line, thin walled, branched, and septate primary hyphae were usually

restricted to epidermal cells in contact with the penetration peg while thin walled, sparingly branched, and irregularly septate se­

condary hyphae began developing in the intercellular spaces near the

point of infection 24-36 hours after inoculation (18).

A number of studies concerning the sporulation of D. poae have

touched on the difficult problem of inducing sporulation on artificial

media (7, 9, 10, 14, 17, 20). A review of this work is unnecessary

since an excellent discussion has been prepared by Morrison (17).

The influence of temperature on conidium production of I), poae has been

addressed by several authors (9, 17, 21). Halisky and Funk (9) have re­

ported that D. poae readily sporulated on naturally infected leaves

collected from November to April but rarely on leaves collected between

May and October. Poor sporulation of D. poae on leaves collected during

the warmer seasons has been attributed to the presence of an unidentified

heat-induced metabolite in host tissue (9). Colbaugh and Beard (2) have stated that sporulation of Helminthosporium spp. rarely occurs on

intact plants in the field. On SAYA agar plugs, Vargas and Wilcoxson

(21) observed that sporulation of D. poae was generally good at 15-18 C

but quite poor above 21 C. Sporulation of D. poae was not observed

above 27 C (21).

Ambient temperature at the time of conidium formation has an In­

fluence on conidium development (17). Conidium length as well as the

number of septatlons in a I), poae conidium are greatest at 8-10 C but 77 as temperatures increased, the length and number of septations decreased

(17, 21). At temperatures above 21 C, atypical swellings of conidio- phore cells and germination of apical conidiophore cells have also been observed (17,21).

Field studies have raised questions concerning the influence of

temperature on Drechslera poae and disease development which have yet to be answered. The objective of this project was to study the in­

fluence of temperature on the behavior of D. poae on leaf surfaces,

penetration and colonization of leaf tissue, lesion enlargement and

sporulation on host tissue.

MATERIALS AND METHODS

Plant cultivation: 'Park' Kentucky bluegrass (Poa pratensis L.) was seeded at a rate of 80 to 100 seeds/pot in 8.8 cm styrofoam cups filled to within 1 cm of the top with a 1 :1:1 steamed soil, sand, and vermiculite potting mix. Pots of plants were maintained in the greenhouse, routinely clipped to 10 to 15 cm and treated with 200 mg/L of a 15-15-15 water soluble fertilizer. Pots of plants were 6-8 months old when used in experiments.

Inoculum: A Isolate of Drechslera poae was obtained from Richard

H. Morrison of Northrup King Co., Eden Prairie, MN. Stock cultures of fungus were maintained on silica gel to insure a consistant source of inoculum (20). Cultures of Drechslera poae was produced on V-8 juice agar at 18 C with 12 hours fluorescent illumination (4 Klux).

Inoculation : Drechslera poae conidia were harvested from 21 day-old cultures by placing the media from several petri plates in a

1-liter flask with 50 ml of double distilled water, plus one drop of

Tween 20 (Emulsion Engineering, Inc., Elk Grove Village, IL). The 78

The suspension was shaken and then strained through one layer of 80 mesh cheese cloth. A Sedgwick-Rafter counting cell (Hausser Scientific,

Blue Bell, PA) was used to adjust the concentration of the conidium suspension to 10,000 conidia/ml. Conidium suspensions were uniformly applied to Kentucky bluegrass foliage with an artist's air brush at.7 kg/cm^ for 10 sec. Inoculated plants were then sealed in plastic bags for Incubation.

Influence of temperature on pre-penetration: Inoculated pots of

Kentucky bluegrass were incubated at 6 , 12, 18, 24 and 30 C with 12- hour fluorescent illumination (4 Klux) in plastic bags which served as a moist chamber. Leaf samples were collected from pots 2, 4, 6 ,

8 , 12, 16, 24, 32 and 48 hours after inoculation. After removal of the leaf samples, the pots were placed back at the same temperature after removal of the plastic bag.

The leaf samples were taped to 15 x 22 mm index cards. Collodion solution (Fisher Scientific Co., Pittsburgh, PA) was applied to the excised leaves with a tapered glass rod and allowed to dry. Dried collodion strips were removed from the leaves with forceps, placed on a glass slide and stained with 0.5% cotton blue in lactophenol.

Germination, germ tube elongation and appressorium formation of 50

D. poae conidia per treatment on collodion strips was determined at

100X and 400X magnification. Conidia were considered germinated when the germ tubes exceeded 2 urn.

Disease severity: Pots were held at the initialIncubation“temperatuVe for 5 days after inoculation. All foliage was removed for disease evalua­ tion by clipping each pot 1 cm above the lip of the styrofoam cup. 79

The Incidence and severity of the leaf spot phase was evaluated

on 25 randomly selected leaves using the Horsfall and Barratt rating

system (11, 12). The Horsfall and Barratt grading system (11, 12) was

based on the percent leaf area diseased and was as follows: 1 - 0%,

2 - 0-3%, 3 - 3-6%, 4 - 6-12%, 5 - 12-25%, 6 - 25-50%, 7 - 50-75%,

8 - 75-87%, 9 - 87-94%, 10 - 94-97%, 11 - 97-100% , 12 - 100%.

Leaf lesion enlargement: *Parkf Kentucky bluegrass leaves were

collected five days after inoculation from pots held in moist chambers

for 16, 24, 36, and 48 hours at 6 , 12, 18, 24, and 30 C. After

several leaves were taped to a glass slide, the length and width of

20 lesions were measured with an eyepiece micrometer at 40X and 100X

magnification. Since many lesions were elliptical in shape, lesion

area was determined using the formula for the area of an ellipse

(A - ab where a - length and b * width).

Penetration and colonization of leaf tissue: Leaf samples were

collected from pots of plants incubated at 6 , 12, 18, 24, and 30 C

after 16, 24, 32, and 48 hours incubation in a moist chamber, steam­

ed in 0.5% trypan blue in lactoglycerine for 10 minutes, destained in lactoglycerine and mounted in lactoglycerine on glass slides for examination*

A second set of leaf samples were collected from all the 24 hour

treatments, steamed in 0.5% cotton blue in lactophenol for 10 minutes, destained in clear lactophenol for 2 minutes, and mounted on glass

slides for examination.

Leaf pieces were examined microscopically for IK poae conidia, appressoria, and the presence of primary or secondary hyphae in host

tissue. The number of successful penetrations was determined using 80 leaf material from the 24 hour treatment by noting the association be­ tween appressoria and the development of primary hyphae in leaf tissue.

Infection efficiency: Pots of 'Park' Kentucky bluegrass which were inoculated with a D. poae conidium suspension were incubated at

6 , 12, 18, 24, and 30 C in sealed plastic bags to provide a moist environment with 12 hour fluorescent illumination (4 Klux). After

24 hours, the plastic bags were removed and the pots were incubated for five days to allow lesion development. Leaf sections of 10-15 cm were collected from each treatment and examined with a diseccting microscope at 30 to 80X magnification. Infection efficiency, i.e.,

the proportion of conidia successfully giving rise to a lesion was expressed as the number of lesions per leaf section divided by

the total number of conidia observed on the section ^8 ) .

Sporulation of D. poae on excised leaves: Diseased leaves were collected from pots of 'Park' Kentucky bluegrass inoculated with a suspension of 10,000 D_. poae conidia/ml, incubated at 18 C in

plastic bags for 24 hours, removed from the plastic bags, and held

another 4 days at 18 C. Excised diseased bluegrass leaves were placed on sterile filter paper in petri dishes and allowed to air dry for

24 hours to halt fungal activity. Double distilled water was added

to remoisten the dried leaves and the petri dishes were sealed with

parafilm to provide a moist environment. The petri plates were in­

cubated at 6 , 12, 18, 24, and 30 C with 12 hour fluorescent illumina­

tion (4 Klux).

Sporulation of D_. poae was monitored after 2,4, and 6 days incu­

bation. Approximately 5-7 leaves were placed in small vials containing

2 ml of 1Z CuSO^ and shaken vigorously to remove conidia from conidio- phores. The leaves were removed from the vials, folded in a piece of aluminum foil, dried at 80 C for 24 hours, and weighed. The con­ centration of conidia in the CUSO4 solution was determined with a

Sedgwick-Rafter counting cell. The number of conidia/rag dry weight of leaf tissue was calculated by dividing the total number of D. poae conidia in the 2 ml CuSO^ solution by the weight (rag) of leaf tissue.

Comparison of D. poae sporulation on intact plants and excised leaves

Pots of 'Park' Kentucky bluegrass were inoculated with a conidium suspension of 10,000 EL poae conidia/ml, incubated 24 hours at 18 C

in plastic bags, and held another 5 days at 18 C. Leaves from half

the pots were harvested, placed in petri plates and dried while the

remaining pots were held an additional day at 18 C. After 24 hours,

the leaves in the plates were remoisfrened and the remaining pots of

Kentucky bluegrass were sealed in plastic bags. Petri plates and

pots of Kentucky bluegrass were incubated six days at 12, 18, and

24 C with 12 hour fluorescent illumination (4 Klux). After six days,

leaf samples were collected from both treatments, placed in vials

containing 2 ml 1% CUSO4 , and the concentration of D. poae conidia was determined using the technique described in the previous section .

Influence of temperature on conidium and conidiophore morphology:

Small leaf pieces were collected from excised Kentucky bluegrass leaves

incubated for six days at 6 , 12, 18, 24 and 30 C. The samples were mounted on glass slides in 0.5% cotton blue in lactophenol. Conidium

length and width at the apical, mid spore, and basal cell, conidiophore

length and width as well as the number of septa in a conidium were measured or determined at 400X magnification. 82

RESULTS

Influence of temperature on conidium germination, germ tube elonga­ tion and appressorium formation of Drechslera poae on Kentucky bluegrass leaves: Conidium germination of IK poae was first observed on Kentucky bluegrass leaves four hours after Inoculation at all temperatures

(Figure 2.1). Initially conidium germination was most rapid at 18 and 24 C, moderate at 30 C, and slow at 6 and 12 C (Figure 2.1). At

18, 24, and 30 C, IK poae conidium germination was nearly completed within 24 hours of inoculation. Approximately 32 hours after inocu­ lation, the percentage of germinated conidia on Kentucky bluegrass leaves reached the 90% level at 6 and 12 C.

Shortly after germination, germ tube development was quite rapid at 18, 24, and 30 C while only modest increases in length were noted at 6 and 12 C (Figure 2.2). The bulk of germ tube development occurred within 24 hours of inoculation at 18, 24, and 30 C with moderate in­ creases being observed in the following 24 hours. At 6 and 12 C, germ tube development although slower than at the other temperatures, steadily increased throughout the 48 hour period following inoculation.

After 48 hours, the mean lengths of D. poae germ tubes formed at 12,

18, and 24 C were significantly greater than the mean lengths of germ tubes developing at 6 and 30 C (Figure 2.2). Appressorium forma­ tion was most rapid at 18 and 24 C within 24 hours of inoculation but only moderate increases in the number of appressoria were noted during the following 24 hour period (Figure 2.3). The rate of appressorium forma­ tion of D. poae initially was slower at 6 , 12, and 30 C than at 18 or 24 C.

However, afteV 48 hours, no substantial differences In appressorium Figure 2.1 Influence of temperature on the germination of Drechslera poae conidia on leaves of 'Park' Kentucky bluegrass. Leaf samples were collected from pots of Kentucky bluegrass incubated at 6 , 12, 18, 24, and 30 C for 2, 4, 6 , 8 , 12, 16, 24, 32, and 48 hours after inoculation, collodion leaf lnpressions were made, stained and mounted. A total of 100 conidia/treatment were checked for the presence of germ tubes. Conidia were con­ sidered germinated when germ tubes exceeded 2 pm in length. % % germination 100 40 20 80 60 4 8 2 16 12 8 6 4 2 Period \ x A f o nuain h) na moist ina environment (hr) incubation 24 / v 30C 0 -3 I2C - 6C - 18 •C 32 ■ 24C 48 85

Figure 2.2 Influence of temperature on the elongation of Drechslera poae germ tubes on leaves of 'Park' Kentucky bluegrass. Leaf samples were collected from pots of Kentucky bluegrass incubated at 6, 12, 18, 24, and 30 C for 2, 4, 6, 8, 12, 16, 24, 32, and 48 hours after inoculation, collodion leaf impressions were made, stained and mounted. The germ tubes of a total of 100 conidia/treat­ ment were measured with an eyepiece micrometer at 10QX and 400X magnification. Mean germ tube length \x.m 120 180 140 160 100 0 4 0 6 20 0 8 2 16 12 8 6 4 2 eid f nuain h) n mos evrn ent environm oist m a in (hr) incubation of Period «• 24C «• I8C - I2C - 6C - 300 0 3 - 2 3 4 2 48 87

Figure 2.3 Influence of temperature on the formation of appressoria of Drechslera poae on leaves of 'Park* Kentucky bluegrass. Leaf samples were collected from pots of Kentucky bluegrass incubated at 6, 12, 18, 24, and 30 C for 2, 4, 6 , 8, 12, 16, 24, 32, and 48 hours after inocu­ lation, collodion leaf impressions were made, stained, and mounted. The germ tubes of 100 conidia/treatment were examined for the presence of appressoria. 2 4 6 8 12 16 2 4 3 2 4 8 Period of incubation (hr) in a moist environment

00 00 89 formation were observed on Kentucky bluegrass leaves at 12 to 30 C.

Only at 6 C were significant reductions in appressorium formation observed.

Influence of temperature on the penetration and colonization of

'Park' Kentucky bluegrass leaves: Incubation temperature had no apparent effect on the site of appressorium formation. Appressorium formation and subsequent penetration of Kentucky bluegrass leaves by IK poae generally occurred at the juncture of the walls of two adjacent epi­ dermal cells. Few penetrations through stomatal aperatures were noted although the formation of appressoria over stomates was not uncommon at any temperature. Rather, the subsidiary cells surrounding the guard cells were penetrated and colonized. Drechslera poae rarely penetrated through the epidermal cells to the intercellular spaces surrounding the palisade layer but colonized epidermal cells prior to movement between the palisade cells.

The number of successful penetrations of Kentucky bluegrass leaves was significantly greater at 18 and 24 C than at 6, 12, and 30 C

(Table 2. 1). However, it must be noted even at 6 and 30 C, nearly

50% of the appressoria on Kentucky bluegrass leaves were associated with infection sites (Table 2, 1).

Successful penetration of epidermal cells was indicated by the forma­ tion of a large globose ceil known as the primary hyphal initial (Figure

2.4 A,B). Shortly after development of the primary hyphae Initial, primary hyphal filaments which were often thin walled, expanded and regularly septate began to grow from the primary hyphal initial (Figure 2.5 A.B.)

Formation of primary hyphal initials and hyphae were commonly associated with appressoria on the leaf surface at 18 and 24 C 90

Table 2.1 Influence of temperature on the penetration of of Kentucky bluegrass leaves by Drechslera poaex .

% successful v Temperature C penetrations'

6 46 b z

12 50 b

18 67 a

24 65 a

30 41 b

Inoculated pots of ’Park' Kentucky bluegrass were incubated 24 hours at 6, 12, 18, 24, and 30 C in a moist environment and held a further four days at the same temperature. Leaves were collected and cleared by steaming the leaves in 0.5% lactophenol cotton blue. Six to eight leaves/treatment were mounted on slides in lactoglycerine. Examination of the leaves focused on the association between appressoria and primary hyphae in leaf tissue. y 'Penetration was successful when primary hyphae began to develop in leaf tissue. z Values in the column followed by the same letter are not significantly different from one another at the P*0.05 level according to Duncan's Multiple Range Test. 91

Figure 2.4 Pre-penetration events of Drechslera poae on leaves of Kentucky bluegrass. A) germinated conidia (c) approximately 24 hours incubation, note extensive germ tube (gt) development and several appressoria(a). B) germinated conidium with short germ tube tipped with an appressorium. C) appressoria sometimes formed over stomates (s) but D) most often at the cell wall juncture of two epidermal cells. 92 93 and uncommon at 12 C approximately 16 hours after inoculation. At

6 and 30 C. penetration and development of hyphal initials were not noted until 24 hours after inoculation. Primary hyphae in epidermal cells continued to expand for several hours prior to movement into the palisade layer. The length of primary hyphae varied from 1 pm to 200 um in some epidermal cells. The rate of elongation of primary hyphae in epidermal cells was more rapid at 12, 18, and 24 C than at 6 or 30

C. Hyphae of 40 um or more were present in epidermal cells within 24 hours when incubated at 12, 18, and 24 C but were not observed in epidermal cells incubated at 6 or 30 C until 32 to 48 hours after inoculation.

Infrequently, appressoria-like swellings on the lateral walls of the colonized epidermal cells were observed developing at 12, 18,

24, and 30 C. A distinct pore was formed in the cell wall between the two cells and primary hyphae began to develop in the newly colo­ nized cell. It was impossible to determine whether the cell was alive when penetrated or what effect penetration had on cellular components.

Development of secondary hyphae in the intercellular spaces of bluegrass rapidly followed primary hyphae formation at 18 and 24 C.

Secondary hyphae filaments which were thin walled, narrow, and irregu­ larly septate, were present 16 hours after inoculation. Extensive ramification by secondary hyphae occurred within 24 to 32 hours of

inoculation. Development of secondary hyphae in leaf tissue at 12 C was not observed until 24 hours after Inoculation. At 6 and 30 C, secondary hyphal development was sparse 48 hours after inoculation.

Direct penetration and colonization of palisade cells by secondary hyphae was not observed. 94

Palisade cells often did not visibly react to the presence of

D. poae hyphae in epidermal cell layer. Lesion development coincided with the ramification of secondary hyphae and plasmolysis of the sur­ rounding cells in the palisade layer (Figure 2.5 C). However, at 6 C, moderate ramification of D. poae secondary hyphae in the palisade layer was not accompanied by lesion formation. Based on lesion size, the ramification of D. poae hyphae was much more extensive at 12, 18, and

24 C than at 6 or 30 C.

Once lesion formation had begun, secondary hyphae were not observed

outside the lesion margin. Lesion expansion was limited by the pre­

sence of vascular bundles in leaf tissue which limited secondary hy­

phal development (Figure 2.5D). Inhibition of secondary hyphal rami­

fication by vascular bundles may cause the formation of characteristic

or oval leaf lesions (Figure 2.5D).

Influence of temperature on the development of the leaf spot phase

of melting-out: Initially, temperature had a substantial effect on the

incidence of the leaf spot phase of melting-out caused by D. poae.

Penetration of leaf tissue and subsequent leaf lesion formation began

within four hours of inoculation at 18 C (Figures 2.6, 2.7). At 12

and 24 C, penetration occurred within eight hours of inoculation (Fig­

ure 2.6). Once penetration began, the leaf spot incidence rapidly

increased to the point where nearly all leaves were diseased at 12, 18,

and 24 C (Figure 2.6). Leaf spot initiation was inhibited at 6 and 30 C

until 12 hours after inoculation (Figure 2.6). As the duration of

incubation of inoculated plants continued, leaf spot incidence greatly

increased. After 32 hours incubation, no substantial differences in

disease incidence were noted between the five treatments (Figure 2.6). 95

Figure 2.5 Penetration and colonization of Kentucky bluegrass leaves by Drechslera poae. A) the primary hyphal initial (phi) forms immediately after penetration in host epidermal cells or intercellular spaces. Primary hyphae (ph) develops from the primary hyphal initial in epidermal cells or intercellular spaces (here) (320X). B) primary hyphal initial and primary hyphae in epidermal cell (1000X). C) secondary hyphae (sh) ramifying between palisade cells (pc) (1000X). D) extensive secondary hyphal development near a vascular bundle (1000X). ■

96 Figure 2.6 Influence of temperature and moisture on the incidence of the leaf spot phase of melting-out of Kentucky bluegrass caused by Drechslera poae- Five days after inoculation, the foliage of each pot was harvested and disease incidence determined. Disease incidence was based on the percentage of leaves with leaf lesions per 200 leaves examined for symptom development. Disease inctd*** 99

Figure 2.7 Influence of temperature and moisture on the severity of the leaf spot phase of melting-out of Kentucky bluegrass caused by Drechslera poae. Five days after inoculation, the foliage of each pot was harvested and disease severity determined. Disease severity ratings were based on the Horsfall and Barratt grading system which is as follows: 1*0%, 2*0-3%, 3*3-6%, 4* 6-12%, 5*12-25%, 6*25-50%, 7=50-75%, 8*75-87%, 9*87- 94%, 10*94-97%, 11*97-100%, 12-100% of the leaf area was diseased. Disease severity 4.0 3.0 5.0 2.0 46 1 1 2 32 24 16 12 8 6 : 4 eid f nuain h) n mit environment moist a in (hr) incubation of Period * 30C 24C I8C

48 100 101

Figure 2.8 Influence of temperature on leaf lesion enlargment in a moist environment. Inoculated pots of 'Park' Kentucky bluegrass were removed from plastic bags 16, 24, 32, and 48 hours after inoculation. Lesion dimensions were measured five days after inoculation and lesion area was calculated using the formula for the area of an ellipse. Lesion area 2.0 _ 3,0 1.0 6 4 2 8 6 24 16 48 32 24 16 6 C 12 1 2 C 24 C 18 C iJL \ v Period (hrs) in a moist environment (hrs)a in moist Period 2 8 6 4 2 8 6 4 2 48 32 24 16 48 32 24 16 48 32 Incubation temperature Incubation m m ii ii i I . 6 4 2 48 32 24 16 18 0 C 30 ■ K ; ■ ■ i N o 103

Figure 2.9 Influence of temperature on the infection efficiency of Drechslera poae on leaves of 'Park' Kentucky bluegrass. Inoculated pots of bluegrass were incu­ bated 24 hours in a moist environment at 6 , 12, 18, 24, and 30 C. Infection efficiency which was cal­ culated five days after inoculation was determined by dividing the number of lesions observed by the total number of germinated conidia in that sample. 6C I2C I8C 24C 30C Incubation temperature 105

Disease severity, which is a logarithmic scale based on the per­ cent area diseased rather than percentage of leaves diseased, provided a different view of disease development. Disease severity increased slower than disease incidence rating reflecting the gradual expansion of lesions on Kentucky bluegrass leaves maintained in a moist environ­ ment (Figure 2.7). Increases in disease severity were comparable at

12, 18 and 24 C until 16 hr then slightly higher at 18 C from 16-48 hr

(Figure 2.7). Lesion development was noticeably inhibited at 6 and 30 C even after 48 hours incubation in a moist environment.

Moisture also plays an important role in leaf lesion formation.

Besides being required for conidium germination and infection, mois­ ture is also necessary for lesion expansion (Figures 2.7, 2.8). Lesion area increases in proportion to the length of the incubation period in a moist environment (Figures 2.7, 2.8). Lesion expansion occurred at approximately the same rate at 12, 18, and 24 C (Figure 2.8). At

6 and 30 C, the low disease severity ratings reflected in the slow rate of increase of leaf lesion area (Figure 2.8).

Influence of temperature on infection efficiency: The efficiency of Infection of bluegrass leaves Infected with D. poae peaked at a temperature of 18 C (Figure 2.9). Although efficiency values were significantly different from one another, infection efficiency values at 12, 24, and 30 C were moderate. Only at 6 C, was lesion develop­ ment effectively suppressed. 106

Sporulation of Drechslera poae on Kentucky bluegrass leaves:

Conldiophore development began on excised leaves after 24-32 hours

Incubation in a moist environment. Sporulatlon of D. poae was

quite light at all temperatures after two days (Figure 2.10). Tre­ mendous increases in the level of D. poae sporulatlon at 1 2 , 18,

and 24 C were observed after four days incubation. This increase

in conidium production closely followed extensive conldiophore de­

velopment on the surface of excised Kentucky bluegrass leaves (Figures

2.10, 2.11). At 6 and 30 C, conldiophore development was confined

to areas within the lesion margin. Sporulation of D. poae at 6 C,

though greatly inhibited, did continue to increase slowly in intensity

while at 30 C sporulation was sparce after six days incubation

(Figure 2.10). Saprophytic activity of Aspergillus and Alternaria spp.

was quite high at 24 C and especially at 30 C. Very few saprophytic

fungi were observed colonizing the excised leaves at 6 , 12, or 18 C.

Sporulation of _D. poae was not as profuse on intact plants

as on excised leaves (Table 2.2). The level of sporulation of I), poae

on excised leaves was 4 times greater at 12 and 18 C to 10 times

greater at 24 C than on leaves from intact plants. Temperature did

not significantly influence sporulatlon on leaves of intact plants

(Table 2.2).

Influence of temperature on the morphological characters of

Drechslera poae conidia and conidiophores: Ambient temperatures had

a substantial influence on the length and number of septae in a

conidium as well as the conldiophore length (Table 2.3,Figure 2.12).

Mean conidium length and number of septate was highest at 12 C and 107

Figure 2.10 Sporulation of Drechslera poae on excised, diseased 'Park' Kentucky bluegrass leaves at five temperatures. Conidium production was monitored after 2, 4, and 6 days incubation at 6 , 12, 18, 24, and 30 C by dividing the number of I), poae conidia washed from leaves by the dry weight of the leaves (mg). g O 18 5 O o fo o o o No. conidia/mgNo. dry leaf weight o > 0

Period of incubation Idaysl 80T 109

Table 2.2 Sporulation of Drechslera poae on diseased, excised leaves and intact 'Park' Kentucky bluegrass plants after six days Incubation in a moist environment at three temperatures.x

# conidia/mg dry leaf weighty Temperature C excised leaves intact plants

12 1408 aZ 280 be

18 1267 a 329 be

24 438 b 48 c

^ots of 'Park' Kentucky bluegrass were inoculated with 10,000 conidia/ml, incubated 24 hours at 18 C, and held a further five days at 18 C. Foliage from half the pots was harvested and dried for 24 hours. Simultaneously, the dried leaves and remaining pots were placed in a moist environment for six days at 12, 18, and 24 C with 12 hours fluorescent il­ lumination. Leaves were collected from both treatment sets, places in vials containing 2 ml of 1% C11SO4 solution, and shaken. The concentration of D. poae conidia in the 1% CUSO4 solution was determined with a Sedgwick-Rafter cell. Leaf material was collected from the vials, dried and weighed.

^The number of conidia/mg dry leaf weight was calculated by dividing the total number of conidia in 2 ml of 1% CUSO4 solution by the dry weight (mg) of the leaves. 2 Values followed by the same letter are not significantly different from one another at the P*0.05 level according to Duncan's Multiple Range Test. 110

Figure 2.11 Profuse sporulatlon of Drechslera poae on a diseased, excised Kentucky bluegrass leaf (80X). HI 112

Figure 2.12 Influence of temperature on Drechslera poae conidium morphology. Note the differences in length and number of septations on these Drechslera poae conidia - collected from excised Kentucky bluegrass leaves after six days incubation at A) 12 C, B) 18 C, C) 2k C, and D) 30 C (all 500X). 113

rfV

CD

" t Table 2.3 Influence of temperature on the morphological characters of Drechslera poae conidia and conldlophores developing on lesions on 'Park* Kentucky bluegrass.2

Conidium Morphological 6 C 12 C 18 C 24 C 30 C Characters______

Length pm Mean 118.9 141.1 122.5 85 50 Range 71-173 103.5-181.7 86.9-156.9 50.6-115 25.3-75.9

Basal septum width pm

Mean 16.8 18.3 17.8 15.3 15.2 Range 12.4-20 11.5-23.5 14.3-23 11.5-18.4 9.2-20.9

Mid-spore width pm Mean 19.7 20.5 20.5 18.3 17.2 Range 17.9-21,9 16,1-23.2 16.6-25.3 14.3-23 13.1-23

Apical septum width pm Mean 15 14.7 14 12 16.8 Range 11.5-19.3 11.5-18.4 11.5-18.9 9.2-14.3 11.7-25.8

Septae (no.) Mean 8.7 10.4 8.6 6 3.2 Range 5-11 7-14 5-12 3-9 1-5 Table 2.3 (cont.) Influence of temperature on the morphological characters of Drechslera poae conldia and conidiophores developing on lesions on 'Park’ Kentucky bluegrass.

Conidiophore Morphological 6 C 12 C 18 C 24 C 30 C Characters

Length pm Mean 86.6 92.3 114.2 186.5 256.8 Range 43.7-161.5 55-144.9 82.8-154.1 135.7-269.1 128.8-384

Width pm Mean 9.1 9.7 9.5 9.5 9.8 Range 6.9-11.3 8.7-11,5 7.2-13.8 8.6-11.5 8.1-13.8

2 Conidiuro and conidiophore dimensions were measured with an eyepiece micrometer at 400X magnification. Each value is the mean of 25 different observations. 116

Figure 2.13 Influence of temperature on Drechslera poae conidium initial formation. A) At 18 C, conidium development is porogenous, i.e., conidium in­ itials develop through pores or a protrusions of the conidiophore wall. B) At 24 C, conidium initails develop as extensions of the conidiophore tip (500X). 117 118 the values of both parameters decreased in magnitude as the temperature increased or decreased (Table 2 Figure 2.12). Substantial differences in conidium width at the apical, mid-spore or basal septum were not observed (Table 2.3). Murogenous conidium development, i.e., conidium initial developing as an extension of the conidiophore tip rather than through pores or as a protrusion in the condiophore wall, was common at 24 C but very unusual at other temperatures (Figure 2.13).

Mean conidiophore length increased as the incubation temperature increased from 6 to 30 C while conidiophore width was unaffected by changes in temperature (Table 2.3). At 30 C, the apical cell of the conidiophore never differentiated to allow conidium formation and con­ tinued to divide and elongate.

DISCUSSION

Results of this study confirm the field observation of several authors that Drechslera poae activity on leaf surfaces, penetration of leaf tissue and subsequent lesion development is greatest at temperatures from 12, to 24 C with peak activity occurring at 18 C (3,4,9,17,21). The level of D. poae conidium germination, germ tube development, and appressorium formation on

Kentucky bluegrass leaves was moderately to severely restricted at

6 , 12, and 30 C for 16 to 24 hours after inoculation (Figures 2.1 -

2.3). Although D. poae conidium germination was noted at all tempera­ tures after 4 hours, substantial differences in the level of germina­ tion were quite evident. Horsfall (10) noted that approximately 98% of the D. poae conidia examined at temperatures as low as 3 C had germ­ inated within 24 hours. In this study that level of germination at 6 and

12 C did not occur until 32 to 48 hours after inoculation (Figure 2.1). 119

The level of 13. poae conidium germination 24 hours after inoculation at temperatures between 18 to 20 C reached the same levels noted by

Horsfall (10) and Mower (18). Germ tube development was nearly complet­ ed within 24 hours of inoculation at 18 to 30 C (Figure 2.2). How­

ever, Mower (18) noted at 24 to 30 C germ tube development was com­

pleted within 12 hours of inoculation. Probably, the method of ob­

servation used to monitor these phenomena account for the difference be­ tween those observations. Although appressorium formation was inhibited at

6 and 12 C, appressorium formation was generally steady for up to 48 hours

after inoculation (Figure 2.3). It i3 difficult to determine whether

the levels of appressorium formation in this study were comparable to

those of Mower's (18) because different methods for monitoring appres­

sorium formation were used.

The sequence of events pertaining to penetration and colonization

of Kentucky bluegrass by D. poae were nearly the same in this study

as those outlined by Mower (18). Mower (18) stated that primary hyphae

did not colonize adjacent cells by the development of intracellular

hyphae. However, formation of appressorium-like structures in colonized

epidermal cells preceded the penetration and colonization of adjacent

cells by primary hyphae at temperatures from 12 to 30 C.

It must be noted prior to further discussion that the study com­

pleted by Mower (18) was done at temperatures between 24 to 30 C,

temperatures generally higher than those encountered in the field when

Kentucky bluegrass is being penetrated and colonized by D. poae. in

this study, I have attempted to observe phenomena at temperatures above and below field optimums. 120

Temperature extremes did reduce the rate of penetration and colonization of Kentucky bluegrass leaves by JD. poae. Development of primary hyphae indicating successful penetration of an epidermal cell was commonly observed at 18 to 24 C approximately 16 hours after inoculation. Mower (18) first noted primary hyphae developing after 18 hours. However, disease incidence data from this study and one completed by Hallsky and Funk (9) indicate that penetration of

Kentucky bluegrass leaves may begin approximately 6 hours rather than

16-18 hours after inoculation. Penetration follows appressorium

formation by several hours rather then 8 to 10 hours at optimum tem­

peratures (18). Primary hyphal development at 12 C was sparse while

at 6 and 30 C development of primary hyphae was severely restricted.

Disease incidence data again indicate penetration of leaves at 6 and

30 C began in 12 - 16 hours of inoculation yet no successful penetra­

tions were observed on the cleared leaf material incubated 16 hours

at either 6 or 30 C (Figure 2.6). Quite possibly, successful pene­

trations at 6 and 30 C were rare after 16 hours but not uncommon 24

hours after inoculation (Table 2.1).

As observed by Mower (18), ramification of secondary hyphal

filaments into the palisade layer began 6-8 hours after colonization

of epidermal cells 12-24 C. Secondary hyphal development was uncommon

at 6 and 30 C.

Data concerning disease severity and lesion expansion is a good

indicator of secondary hyphae development in host tissue (Figures

2.8, 2.9). Development of primary hyphal filaments often stopped

after several cell divisions. Lesion expansion at 12 to 24 C indica­

tes that ramification of hyphae in the palisade layer was quite rapid. 121

Hyphae of D. poae may be quite sensitive to moisture levels in host tissue. Lesion expansion on intact plants continued even at optimum temperatures for D. poae only if leaves are held in a moist environment. If the leaves were removed from the moist environment, lesion expansion and presumably hyphal growth ceased (Figures 2,8, 2.9).

A similar effect was noted by Halisky and Funk (9).

The infection efficiency of temperature on infection has not been widely studied. Work has concentrated on the influence of inoculum levels on lesion development (5, 8, 19, 22). Peak efficiency occurred at 18 C and decreased significantly with increases or decreases with temperature. At 18 C, the peak efficiency rating of D. poae on Ken­ tucky bluegrass was slightly larger than that of Bipolaris maydis

(Helminthosporium maydis) on com. Infection efficiency of other

fungi such as Septoria lycopersici and Alternaria solani on tomato were much lower than the values for either I), poae or _B. maydis

(15). Given losses due to dispersal and germination, there was a 25-30%.

chance that a conidium will penetrate and produce a lesion under op­

timal conditions (22).

Conidiophore and conidium formation of D. poae on diseased, ex­

cised Kentucky bluegrass plants began approximately 24 to 36 hours at

18 C in a moist environment. Mower (18) reported that conidiophore

and conidium formation began five days after Inoculation of excised

leaves with Ih poae conidium suspension. In Mower's (18) study,

inoculated leaves were incubated continuously in a moist environment while the leaf material used in this study was held in a moist environ­ ment only 24 hours at 18 C to allow penetration and colonization, held for days to permit symptom development, and then remoistened for 122

Incubation.

Temperature has a tremendous Impact on the level of I), poae sporulation on excised leaves of intact plants (Figure 2,10). Sporu- lation occurred most rapidly at temperatures between 12-24 C with peak conidium production occurring at 18 G (Figure 2.10). At 6 and

30 C, conidium production was light and sparce, respectively, after six days incubation on excised leaves (Figure 2.10). Vargas and Wil- coxson (21) noted a different pattern of D. poae sporulation on SAYA agar. Although 18 C was recognized as the temperature optimum for sporulation in both studies, Vargas and Wilcoxson (21) noted tre­ mendous drops in the level of sporulation at 15 and 21 C. On excised

leaves, sporulation of D. poae was profuse from 12 to 24 C, a much

wider range than indicated by Vargas and Wilcoxson (21).

It was difficult to pinpoint the maximum temperature for I), poae

sporulation. In this study, the level of sporulation at 24 C did

vary considerably on excised leaves (Figure 2.10, Table 2.2). Evi­

dently, D. poae was quite sensitive to temperatures between 21 to 24 C

but it was difficult to predict how it will respond to these tempera­

tures at any time.

Conidium production on intact plants was 4 to 10 times lower

than on excised leaves depending on the incubation temperature (Table

2.2). At 12 to 24 C, conidiophores developed over the entire surface

of excised Kentucky bluegrass leaves within four to six days indica­

ting the rapid colonization of the leaf by D. poae. Colonization of

leaves on intact plants appeared to progress at a slower rate. After

six days, approximately 50Z of the area of the leaves collected from

Kentucky bluegrass plants was healthy. The level of sporulation on 123

Murogenous B. soroklniana conidia also formed at temperatures above optimum for conidium formation (13).

Abnormal development of D. poae conidlophores has been observed in several studies (18,21). Twisting and curling as well as germina­ tion of conidlophores has been noted at temperatures above 21 to 24 C

(18). Conidiophore length also increased slgnficantly as temperature increased to 20 C (Table 2.3). It was quite possible that the apical cell of the condiophore was no longer sensitive to wavelength of radiation greater than 480 mm which directed its differentation and initiation of conidium formation (21). LITERATURE CITED

1. BEAN, G. A., and R. D. WILCOXSON. 1964. Pathogenicity of three species of Helminthosporium on roots of bluegrass. Phytopathology. 54:1084-1085.

2. COLBAUGH, P. F. and J. B. BEARD. 1980. Influences of management practices on conidial production by Helminthosporium spp. on Improved Bermudagrass. In P. 0. Larsen and B. G. Joyner, eds. Advances in Turf Pathology, Harvest Publications, Cleveland, Ohio (in press).

3. COUCH, H. B. 1957. Melting-out of Kentucky bluegrass, its cause and control. Golf Course Reptr. 25:5-7.

4. COUCH, H. B. 1973. Diseases of turfgrass. Krieger Publishing Co., Huntington, N.Y. 348p.

5. DAVISON, A. D., and E. K. VAUGHAN. 1964. Effect of uredospore concentration in determination of races of Uromyces phaseoli var. phaseoli. Phytopathology, 54:336-337.

6 . DRECHSLER, C. 1922. A new leaf spot of Kentucky bluegrass caused by an undescribed species of Helminthosporium. (Abstr.) Phytopathology. 13:35.

7. DRECHSLER, C. 1930. Leaf spot and footrot of Kentucky bluegrass caused by Helminthosporium vagans. J. Agr. Res. 40:447-456.

8 . GREGORY, P. H. 1961. The microbiology of the atmosphere. Leonard Hill, London. 25lp. *

9. HALISKY, P. H. and C. R. FUNK. 1966. Environmental factors affect­ ing growth and sporulation of Helminthosporium vagans and its pathogenicity to Poa pratensis. Phytopathology. 56:1294-1296,

10. H0RSFALL, J. G. 1930. A study of meadow-crop diseases in New York. Cornell Univ. Expt. Sta. Mem. 120. 139p.

11. HORSFALL, J. G. 1945. Fungicides and their action. Chronica Botanlca Co., Watham, Mass. 239p.

12. HORSFALL, J. G., and R. W. BARRATT. 1945. An improved grading system for measuring plant diseases (Abstr.) Phytopathology. 35:655.

13. JOHNSON, T. W., and J. E. HALPIN. 1954. Environmental effects on conidial variation in some Fungi Imperfecti. J. Elisha Mitchell Sci. Soc. 70:314-326.

124 125

14. LUKENS, R. J. 1960. Conidial production from filter paper cultures of Helminthosporium vagans or Altemaria solanl. Phytopathology. 50:867-868.

15. McCALLAN, S. E. A., and R. H. WELLMAN. 1943. A greenhouse method of evaluating fungicides by means of tomato foliage diseases. Contr. Boyee Thompson Inst. 13:93-134.

16. MONTEITH, J. 1924. The leafspot disease of bluegrass. U.S. Golf Assoc. Green Sect. Bull. 4:172-173.

17. MORRISON, R. H. 1980. Leaf spot of Kentucky bluegrass in Minne­ sota: Misidentification of Drechslera poae as IK dictyoldes. In P. 0. Larsen and B. G. Joyner, eds. Advances in Turf Pathology. Harvest Publications, Cleveland, Ohio (in press).

18. MOWER, R. G. 1961. Histological studies of suscept-pathogen re­ lationships of Helminthosporium sativum. Helminthosporium va­ gans and Curvularia lunata on leaves of 'Merion' and common Kentucky bluegrass (Poa pratensis L.). Ph.D. Thesis, Cornell Univ. 150p. 19. PETERSON, L. J. 1959. Relations between inoculum density and infection of wheat uredospores of Puccinia graminis var. tritici. Phytopathology. 49:607-614.

20. SLEESMAN, J., P. 0. LARSEN, and J. SAFF0RD. 1974. Maintenance of stock cultures of Helminthosporium maydis (race T and 0). Plant Dis. Reptr. 58:334-336.

21. VARGAS, J. M . , and R. D, WILCOXSON. 1969. Some effects of temperature and radiation on sporulation by Helminthosporium dictyoides on agar media. Phytopathology, 59:1706-1712.

22. WAGGONER, P. E., J. G. HORSFALL, and R. J. LUKENS. 1972. Epimay: a simulation of southern corn leaf blight. Bull. Conn. Expt. Sta., New Haven. 84p. Chapter III

BIOASSAY OF FUNGICIDE RESIDUE PERSISTANCE ON KENTUCKY BLUEGRASS LEAVES ABSTRACT

Iprodione, anilazine, and cyclohexlmide were applied to 1.5 in'1 replicated field plots containing a blend of 'Park* and 'Delta' Ken­ tucky bluegrass. Plugs of turf 12 cm in diameter were removed from each treatment plot 0, 1, 3, 5, 7, 14, and 21 days after fungicide application. Plugs were inoculated uniformly with a suspension of

10,000 Drechslera poae conidia/ml and incubated at 18 C for 24 hours.

At that time, conidium germination, germ tube elongation, and appres­ sorium formation of I). poae on leaves of Kentucky bluegrass were evaluated using the collodion leaf impression technique. Initially, all fungicides effectively suppressed the pre-penetration events of

S.* Poae. Residues of iprodione and anilazine effectively inhibited

D. poae on Kentucky bluegrass leaves for three weeks while little evi­ dence of cyclohyximide residue was noted seven days after fungicide application. The collodion leaf impression technique proved to be a simple means of assaying fungicide residue activity on leaf surfaces under field conditions.

INTRODUCTION

Weather variables such as rain, wind, temperature, and light play' a significant role in the inactivation of fungicide deposits on plant surfaces (5). Greenhouse tests employing artificial weathering agents such as simulated rainfall have proved useful for the evaluation of fungicide persistance however poor correlations between greenhouse and field data may occur (13). Bioassays have been specifically developed to monitor fungicide weathering from foliage in the field. Neely (11)

127 128 evaluated fungicide activity by placing a cellophane disk over a leaf disk collected from fungicide treated plants, pipetting a suspension of Monillnia fructicola on the cellophane disk, and microscopically examining each cellophane disk for germinated conidia. Ko et al (8 ) collected an inoculated treated leaves with Alternaria altemata conidia with a vertical illumination microscope.

Hagan and Larsen (6 ) used a collodion technique that is a simple and useful method for evaluating the effect of fungicides on the conidium germination, germ tube elongation, and appressorium formation of B. sorokiniana on Kentucky bluegrass leaves. The major advantages of the collodion technique over the bioassay described above are that the fungal thallus is permanently mounted for later observation and no special equipment is required.

Melting-out caused by Drechslera poae (Baudys) Shoem. (■Helminthos­ porium poae Baudys, Helminthosporium vagans (Drechs.) Shoem.). is a serious disease wherever Kentucky bluegrass (Poa pratensis L.) is grown in the United States (1,3,7,9,10). Little is known of the direct toxic effects of registered fungicides on D. poae pre-penetration events on leaf surfaces or the rate of fungicide deposit degradation on Kentucky bluegrass leaf surfaces. Generally, fungicide efficacy is evaluated

on the basis of inhibition of symptom development.

The objective of this study was to determine the usefulness of

the collodion technique as a method for evaluating the weathering

of fungicide deposits on Kentucky bluegrass under field conditions

by monitoring conidium germination, germ tube elongation, and appres­

sorium formation of D. poae. 129

MATERIALS AND METHODS

Fungicide application - Iprodione (Chipco 26019, 50WP, Rhone- poulenc, Inc., Mommouth Junction, NJ) anilazine (Dyrene, 50WP, Mobay

Chemical Corp., Kansas City, Mo), and cycloheximide (Acti-dione, 2.1 WP,

Upjohn Co., Kalamazoo, Mich.) were applied at two label rates at a dilution rate of 11.4 L of water/93m^ (3 gal/1000 ft^) using a CO2-

powered sprayer at 2.1 Kg/cm^ to 1.5 m^ replicated and randomized field

plots containing a blend of 'Park' and 'Delta' Kentucky bluegrass on

October 3, 1979. Twelve cm diameter plugs were collected from each

replicate plot 9, 1, 3, 5, 7, 14, and 21 days after fungicide applica­

tion.

Inoculation - An isolate of Drechslera poae was obtained from

Richard H. Morrison of Northrup King Co., Eden Prairie, MN. Stock

cultures of D. poae were maintained on silica gel to insure a con­

sistent source of inoculum (11). Abundant sporulation was induced by culturing IK poae on V-8 juice agar at 18 C with 12 hours fluores­

cent illumination (4 Klux).

Conidia were harvested from 21 day old cultures by placing the

media from petri plates in a 1-liter flask with 50 ml of double dis­

tilled water plus one drop of Tween 20 (Emulsion Engineering, Inc.,

Elk Grove Village, IL), vigorously shaking the flask, and straining

through one layer of cheese cloth. The conidial adjustment of the

concentration in the filtrate was adjusted 10,000 conldla/ml with

a Sedgwick-Rafter counting cell using a Howard eyepiece micrometer.

The conidial suspensions were uniformly applied to the Kentucky

bluegrass foliage with an artist's air brush at .7 kg/cm^ for 10 130

sec. Inoculated plugs were sealed In plastic bags and incubated at

18C with 12 hr. fluorescent illumination (A Klux).

After 2A hours incubation, the plastic bags were removed and leaf

samples were collected from each treatment. Four leaves from each

treatment were taped to 15 x 22 cm index cards. Collodion solution

(Fisher Scientific Co., Pittsburgh, PA) was applied to the leaves with

a tapered glass rod and allowed to dry. Dried collodion strips were

removed from the leaves with a pair of forceps, placed on glass slides

and strained with 0.5% cotton blue in lactophenol for microscopic

examination.

Fifty randomly selected conidia on the collodion strips of each

treatment were evaluated for conidium germination, germ tube elongation

and appressorium formation at 100X and A00X magnification. Conidia were considered germinated when the germ tubes exceeded 2 urn.

Meterological data - Rainfall and turf microclimate temperature

data were collected at a site approximately 50 m from the field plot with a tipping bucket rain gauge and T-type thermocouples, respectively.

A summary of the meterological data is contained in Figure 3.1.

RESULTS

The level of inhibition of Drechslera poae conidium germination, germ tube elongation, and appressorium formation on Kentucky bluegrass leaves provided by iprodione and anilazine which was greater than that provided by cycloheximide completely inhibited conidium germination, germ tube elongation and appressorium formation of D. poae on leaves treated with each fungicide (Figures 3.2 - 3.A). Cycloheximide efficacy decreased at a much faster rate than that observed for either iprodione or anilazine (Figures 3.2 - 3.A). Within seven days of fungicide application, Figure 3.1 Maximum and minimum daily turf microclimate temp­ erature and daily rainfall from October 3-24, 1979 at the OSU Turf Plots, Columbus, Ohio. Temperature C Rainfall (ran) 0 - 30. 20 0 _ 10. 6 4. . L 5. 2 1 . . . . . _ * * * * * ***** 4 6 8 1 1 1 1 1 1 1 1 1 1 2 1 2 3 24 23 22 21 20 19 18 17 16 15 14 13 12 11 10 9 8 7 6 5 4 3 aiu al temperature daily maximum iiu al temperature daily minimum

October ■ ■ ■ ae sampled date * 132 133

Figure 3.2 Effect of fungicide residues on Kentucky bluegrass leaves on Drechslera poae conidium germination. Fungicides were applied to 1.5 m2 replicated field plots of Kentucky bluegrass on October 3, 1979. Plugs were collected 1, 3, 5, 7, 14, and 21 days after fungicide application, inoculated with a Drechslera poae conidium suspension and incubated 24 hours at 18 C. Pre-penetration events were monitored using the collodion leaf impression technique (m*slope of the regression line).

4 % Germination 100 90. 40. 80. 13 1 0 Days between fungicide application fungicide between Days 5 Cycloheximide 7 and inoculation and Iprodione 14 Control Anilazine

21 134 135

Figure 3.3 Effect of fungicide residues on Kentucky bluegrass leaves on Drechslera poae germ tube elongation. Fungicides were applied to 1.5 replicated field plots of Kentucky bluegrass on October 3, 1979. Plugs were collected 1, 3, 5, 7, 14, and 21 days after fungicide application, inoculated 24 hours at 18 C. Pre-penetration events were monitored using the collodion leaf impression technique (m*slope of the regression line). Mean germ tube length pm 100 110 120 40 30. 60. 50. 70. . 3 01 Days between fungicide application fungicide between Days 5 7 n Inoculation and Cycloheximide Iprodlone Anilazine 14 Control

21 136 137

Figure 3.4 Effect of fungicide residues on Kentucky bluegrass leaves on Drechlsera poae appressorium formation. Fungicides were applied to 1.5 replicated field plots of Kentucky bluegrass on October 3, 1979. Plugs were collected 1, 3, 5, 7, 14, and 21 days after fungicide application, inoculated with a Drechslera poae conidium suspension and incubated 24 hours at 18 C. Pre-penetration events were monitored using the collodion leaf impression technique (m“slope of regression line). of Appressoria 10 20 AO. 50. 60. 30. . , 0

1 5 3 Days between fungicide application fungicide between Days 7 and inoculation and Cycloheximide Control 14 Anilazine Iprodione

21 138 Table 3.1 Correlation between the time from fungicide application to inouclatlon with the pre-penetration events of Drechslera poae on Kentucky bluegrass leaves treated with fungicides.

Correlation Coefficient (r) Rate * Conidlum Germ tube Appressorium Treatment (g a.i./93m ) germination elongation formation

Iprodione 56 ,83**Z .56** .61**

Anllazine 112 .77** .51** .67**

Cycloheximide 1.2 .63** .65** .60** * Control - .11 .25 .19

Asterisks: **=statistically significant at the P=0.01 level. 140 no significant differences in conidium germination, germ tube elonga­ tion or appressorium formation of D. poae were noted between the control and cycloheximide treatment values whereas these processes were inhibited below control levels for at least three weeks for iprodione and anilazine (Figures 3.2 - 3.4).

Although well below control values, levels of conidium germination, germ tube elongation, and appressorium formation of D^. poae on leaves treated with each fungicide increased significantly over three weeks

(Table 3.1). The rate of increase of conidium germination, germ tube elongation, and appressorium formation was much faster on leaves treated with cycloheximide than those treated with either iprodione or anilazine (Figures 3.2 - 3.4). In general , the activity of D. poae in­ creased at a slightly faster rate on the iprodione treatments than on the anilazine treatments (Figures 3.2 - 3.4). Since no significant dif­ ferences were noted between the results of the two rates of each fungi­ cide evaluated, the data of the lower rate of each fungicide has been excluded from all tables and figures.

DISCUSSION

The collodion technique proved to be a simple method for evaluating activity of fungicide residues on leaf surfaces of Kentucky bluegrass.

Evaluation of fungicide activity is based on the ability of fungicidal residues on the leaf surface to inhibit the pre-penetration events of

I). poae, conidium gemination, g e m tube elongation, and appressorium formation. Similar tests of fungicide activity have monitored only conidium gemination (8, 11). Evaluation of residual activity of a fungicide based solely on conidium germination may be sufficient for 141

most materials, but some fungicides will selectively inhibit germ tube development or appressorium formation while having little influence on conidium germination (4, 6). Aside from permitting the evaluation of several phenomena on the leaf surface, the easily mounted collodion strips provide a permanent record of fungicide performance. The versa­ tility of this technique allows the simultaneous evaluation of several different fungicides at different rates outdoors.

Apparently, routine mowing and weather variables such as rainfall did not greatly affect persistance of iprodione and anilazine of Ken­ tucky bluegrass leaves. Initially, cycloheximide performed nearly as well as iprodione and anilazine in controlling pre-penetration events of D. poae (Figures 3.2 - 3.4). However, within several days of fungicide application, significant reductions in cycloheximide residue activity were noted (Table 3.1). It was not determined whether the cycloheximide residues were photochemically inactivated or washed from leaf surfaces by rain (Figure 3.1).

The fungitoxicity of iprodione and cycloheximide to Drechslera poae and Bipolaris sorokiniana varies on Kentucky bluegrass leaves (6). Both materials provided excellent control of all D.. poae pre-penetration events immediately after application of either fungicide while neither material effectively suppressed II. sorokiniana conidium germination (6). Anila­ zine suppressed all pre-penetration activity of D. poae and B. sorokiniana on Kentucky bluegrass leaves (6). LITERATURE CITED

1. COUCH, H. B. 1973. Diseases of turfgrass. Krieger Publishing Co., Huntington, N.Y. 348p.

2. DIEKER, U. L. 1955. Sporulation in pure culture by Stemphylium solanj. Phytopathology. 45:141-145.

3. DRECHSLER, C. 1930. Leaf spot and foot rot of Kentucky bluegrass caused by Helminthosporium vagans. J. Agr. Res. 40:447-451.

4. DUNKEL, L. D., R. MAKESHWARI, and P. J. ALLEN. 1969. Structures from rust uredospores: Effect of RNA and protein synthesis inhibitors. Science 163:481-482.

5. EBLING, W. 1963. Analysis of basic processes involved in the deposition, degredation, persistance, and effectiveness of pesticides. Residue Rev. 3:35-163.

6 . HAGAN, A. K. , and P. 0. LARSEN. 1979. Effect of fungicides on conidium germination, germ tube elongation, and appressorium formation by Bipolaris sorokiniana on Kentucky bluegrass- Plant Dis. Reptr. 63:474-478.

7. HALISKY, P. M. and C. R. FUNK. 1966. Environmental factors af­ fecting growth and sporulation of Helminthsoorium vagans and its pathogenicity to Poa pratensis. Phytopathology. 56:1294-1296.

8 . K0, W. H., H. H. HIN, and R. K. KUNM0T0. 1975. A simple method for determining efficacy and weatherability of fungicides on the foliage. Phytopathology. 65:1023-1025.

9. MORRISON, R. H. 1980. Leafspot of Kentucky bluegrass in Minnesota: Misidentiflcation of Drechslera poae as D. dictyoides. In: P. 0. Larsen and B. Joyner, eds. Advances in Turf Pathology. Harvest Publications, Cleveland (In press).

10. MOWER, R. G. 1961. Histological studies of suscept-pathogen rela­ tionships of Helminthosporium sativum, Helminthosporium vagans. and Curvularia lunata on leaves of Merion and Common Kentucky bluegrass (Poa pratensis L.) Ph.D Thesis, Cornell Univ. N.Y. 150p.

11. NEELY, D. 1970. Persistance of foliar protective fungicides. Phytopathology. 60:1583-1586.

12. SLEESMAN, J., P. 0. LARSEN, and J. SAFFORD. 1974. Maintenance of stock cultures of Helminthosporium maydes (race Tand 0). Plant Dis. Reptr, 58:334-336.

142 143

13. TORGENSON, D, C. 1969. Determination and measurement of fungitoxi- city. Pages 93-123 in: D. C. Torgenson, ed. Fungicides, an advanced treatise. Academic Press. London and New York. Vol.I. 697p. BIBLIOGRAPHY

AYLOR, D. E. 1975. Resuspension of particles from plant surfaces by wind. Pages 791-812 in: The Symposium on the Atmosphrere- Surface exchange of particles and gases; Richland, Washington. U.S. Atomic Energy Commission Doc. No. C0NF-740921.

AYLOR, D. E., and P. R. DAY. 1976. Conidium release in Helminthos­ porium (letter to editor).■ Phytopathology. 66:537.

AYLOR, D. E., and R. L. LUKENS. 1974. Liberation of Helminthosporium maydis spores by wind in the field. Phytopathology. 64:1136-1138.

AYLOR, D. E., and J. Y. PARLANGE. 1975. Ventilation required to en­ train small particles from leaves. Plant Physiol. 56:97-99.

BEAN, G. A., and R. D. WILCOXSON. 1964. Helminthosporium leaf spot of bluegrass. Phytopathology. 54:1065-1070.

BEAN, G. A. and R. D. WILCOXSON. 1964. Pathogenicity of three species of Helminthosporium on roots of bluegrass. Phytopathology. 54:1084-1085.

BEARD, J. B. 1973. Turfgrass: science and culture. Prentice-Hall, Inc., Englewood Cliffs, N.J. 658p,

BURCHFIELD, H. P., and A. GOENAGA. 1957. Equipment for producing simulated rain for measuring the tenacity of spray deposits of foliage. Contrib, Boyce Thompson Inst. 19:133-140.

C0LBAUGH, P. F., and J. B. BEARD. 1980. Influence of management prac­ tices on conidial production by Helminthosporium spp. on improved bermudagrass. In P. 0. Larsen and B. G. Joyner, eds. Advances in Turfgrass Pathology. Harvest Publications, Cleveland, Ohio (in press).

COLBAUGH, P. F., and R. M. ENDO. 1974. Drought stress: an Important factor stimulating the development of Helminthosporium sativum on Kentucky bluegrass. Pages 328-333 in: E. C. Roberts, ed. Proc. 2nd Internal. Turfgrass Res. Conf. Amer. Soc. Agronomy.

COUCH, H. B. 1957. Melting-out of Kentucky bluegrass, its cause and control. Golf Course Reptr. 25:5-7.

COUCH, H. B. 1973. Diseases of turfgrasses. Krleger Publishing Co., Huntington, N.Y. 348p. 144 145

DAVISON, A. D., and E. K. VAUGHAN. 1964. Effect of uredospore con­ centration in the determination on races of Uromyces phaseoli var. phaseoli. Phytopathology. 54:336-337.

DIENER, U. L. 1955. Sporulation in pure culture by Stemphylium solanl. Phytopathology. 45:141-145.

DRECHSLER, C. 1922. A new leaf spot of Kentucky bluegrass caused by an undescribed species of Helminthosporium. (Abstr.). Phyto­ pathology. 13:35.

DRECHSLER, C. 1930. Leaf spot and foot rot of Kentucky bluegrass caused by Helminthosporium vagans. J. Agr. Res. 40:447-451.

DUNKEL, L. D., R. MAKESHWARI, and P. J. ALLEN. 1969. Infection struc­ tures from Rust Uredospores: effect of RNA and protein synthesis inhibitors. Science. 163:481-482.

EBL1NG, W. 1963. Analysis of the basic processes involved in the deposition, degredation, persistance, and effectiveness of pesti­ cides. Residue Rev. 3:35-163.

ELLIOT, R. S. 1962. Disease damage on forage grasses. Phytopathology. 52:448-451.

ELLIOT, R. S. 1962. The effect of soil fertility in the development of Kentucky bluegrass diseases. (Abstr.). Phytopathology. 52:1218.

EVERSMEYER, M. G. , C. L. KRAMER, and T. I. COLLINS. 1976. Three suction-type spore samplers compared. Phytopathology. 66:62-64.

GIBBS, A. F., and R. D. WILCOXSON. 1972. Effect of sugar content of Poa pratensis on Helminthosporium leaf spot. Physiol. Plant Pathology. 2:279-287.

GIBBS, a . F., and R, D. WILCOXSON. 1973. The effect of nitrogen fertili­ zation and mowing on Helminthosporium leaf spot and sugar content of bluegrass leaves. Plant Dis. Reptr, 57:544-548.

GRAY, P. M., and J. W. GUTHRIE. 1977. The Influence of sprinkler ir­ rigation and postharvest residue removal practices on the seedborne population of Drechslera poae on Poa pratensis 'Merlon’. Plant Dis. Reptr. 61:90-93.

GREGORY, P. H. 1961. The microbiology of the atmosphere. Leonard Hill, London. 251p.

HAGAN, A. 1980. Isolation, identification, and taxonomy of Drechslera and Bipolarls species on turfgrasses. In: P. 0. Larsen and B. G. Joyner, eds., Advances in Turfgrass Pathology. Harvest Publication, Cleveland, Ohio. 146

HAGAN, A., and P. 0. LARSEN. 1979. Research on Helminthosporium melting- out of Kentucky bluegrass. Proc. Ohio Turfgrass Conf. 1979:46-54.

HAGAN, A., and P. 0. LARSEN. 1979. Effect of fungicides on conidium germination, germ tube elongation, and appressorium formation by Bipolaris sorokiniana on Kentucky bluegrass leaves and disease incidence. Plant Dis. Reptr. 63:474-478.

HALISKY, P. M . , and C. R. FUNK. 1966. Environmental factors affecting growth and sporulation of Helminthosporium vagans and its pathogenicity to Poa pratensis. Phytopathology. 56:1294-1296.

HALISKY, P. M., C. R. FUNK, and R. E. ENGEL. 1966. Melting-out of Kentucky bluegrass varieties by Helminthosporium vagans influenced by management practices. Plant Dis. Reptr. 50:703-706.

HAMILTON, J. M. 1959. Evaluation of fungicides in the greenhouse. Pages 253-257 in: C. S. Holton et^ al^ eds. Plant Pathology: problems and progress, 1908-1958. University of Wisconsin Press, Madison. 588p.

HIRST, J. M. 1952. An automatic volumetric spore trap. Ann. Appl. Biol. 39:257-265.

HIRST, J. M. 1959. Spore liberation and dispersal. Pages 529-538. in: C. S. Holton et al, eds. Plant Pathology: problems and pro­ gress, 1908-1958. University of Wisconsin Press, Madison. 588p.

HIRST, J. M. , and 0. J. STEDMAN. 1963. Dry liberation of fungus spores by raindrops. J. Gen. Microbiol. 33:335-344.

HODGES, C. F. , and W. M. BLAINE. 1974. Midsummer occurrence of Helmin­ thosporium sorokinianum leaf spot on selected cultivars of Poa pra- tensis. Plant Dis. Reptr. 58:448-450.

H0RSFALL, J. G. 1930. A study of meadow crop diseases in New York. Cornell Univ. AGr. Expt. Sta. Mem. 130. 139p.

HORSFALL, J. G. 1945. Fungicides and their action. Chronica Botanica Co., Watham, Mass. 239p.

HORSFALL, J. G., and R. tf. BARRATT. 1945. An Improved grading system for measuring plant diseases. (Abstr.). Phytopathology. 35:655.

INGOLD, C. T. 1970. Fungal spores, their liberation and dispersal. Clarendon Press, Oxford, England. 302p.

INGOLD, C. T. 1971. Liberation mechanisms of fungi. Pages 102-115 in: P. H. Gregory, and J. L. Monteith, eds. Airborne Microbes, 17th Sympos­ ium of the Society for General Microbiology. University Press, Cam­ bridge, fngland. 385p. 147

JAMES, W. R. 1962. The dispersal of spores of Botrytls clnerea Fr. in a raspberry plantation. Trans. BrI> . raycol. Soc. 45:549-559.

JOHNSON, T. W,, and J. E. HALPIN. 1954. Environmental effects on conidial variation in some Fungi Imperfect!. J. Elisha Mitchell Sci. Soc. 70:314-326.

KENNETH, R. 1964. Conidial release in Helmlnthosporla. Nature. 202-1025-26.

KO, W. H., H. LIN, and R. K. KUNIM0T0. 1975. A simple method for determining efficacy and weatherability of fungicides on foliage. Phytopathology. 65:1023-1025.

KRAMER, C. L., M. G. EVERSMEYER, and T. I. COLLINS. 1976. A new 7- day spore sampler. Phytopathology. 66:60-61.

KRAMER, C. L., and S. M. PADY. 1966. A new 24-hour spore sampler. Phytopathology. 56:517-520.

LANGENBERG, W. J., J. C. SUTTON, and T. J. GILLESPIE. 1977. Relation of weather variables andperiodicities of airborne spores of Alternaria dauci and other fungi. Phytopathology. 67:879-883.

LEACH, C. M. 1975. Influence of relative humidity and re-infrared radiation on violent spore release by Drechslera turcica and other fungi. Phytopathology. 65:1303-1312.

LEACH. C. M. 1976. Further comments on spore release by Drechslera (Helminthosporium) spp. and other fungi. (letter to editor) Phytopathology. 66:1265-1266.

LEACH, C. M, 1976. An electrostatic theory to explain violent spore liberation by Drechslera turcica and other fungi. Mycologia. 68:63-86.

LEACH, C. M. 1980. An apparatus for the precise control of humidity, temperature, airflow, and light in spore discharge studies. Phyto­ pathology. 70:189-191.

LEACH, C. M. 1980. Influence on humidity and red-infrared radiation on spore discharge by Drechslera turcica-additional evidence. Phyto­ pathology. 70:192-196.

LEACH, C. M. 1980. Evidence for an electrostatic mechanism in spore discharge by Drechslera turcica. Phytopathology. 70:206-213.

LEACH, C. M., R. A. FULLERTON, and K. YOUNG. 1977. Northern leaf blight of maize in New Zealand. Relationship of Drechslera turcica airspora to factors influencing sporulation, conidium development, and chlamy- dospore formation. Phytopathology. 67:629-636. 148

LEDINGHAM, R. J., and S. H. F. CHINN. 1955. A floatation method for obtaining spores of Helminthosporium sativum from soil. Can. J. Bot. 33:209-303.

LEFEBVRE, C. L., F. L. HOWARD, and F. V. GRAU. 1953. How to keep turfgrass healthy. Pages 285-291. in: Plant Diseases, The Year­ book of Agriculture. U.S. Dept, of Agr., Washington, D.C. 939p.

LUKENS, R. J. 1960. Conidia production from filter paper cultures of Helminthosporium vagans and Alternaria solani. Phytopathology. 50:867-868.

LUKENS, R. J. 1968. Low light intensity promotes melting-out of bluegrass. (Abstr.). Phytopathology. 58:1058.

LUKENS, R. J. 1970. Melting-out of Kentucky bluegrass, a low sugar disease. Phytopathology. 60:1276-1278.

McCALLAN, S. E. A. 1957. Effectiveness of various formulations of Bordeaux mixture in controlling early and late blights of tomato in greenhouse tests. Contrib. Boyce Thompson Inst. 19:157-167.

McCALLAN, S. E. A., and R. H. WELLMAN, 1943. A greenhouse method of evaluating fungicides by means of tomato foliage diseases. Contrib. Boyce Thompson Inst. 13:93-134,

MELCHING, J. S. 1974. A portable self-contained system for the continuous electronic recording of moisture conditions on the surfaces of living plants. ARS-NE-42, Agricultural Research Science, USDA, Washington, D.C. 13p.

MEREDITH, D. 1963. Violent spore release in some Fungi Imperfecti. Ann. Bot. 27:39-47.

MEREDITH, D. 1963. Further observations on the zonate eyespot fungus, Drechslera gigantea in Jamaica. Trans. Brit, raycol. Soc. 46:201-207.

MEREDITH, D. 1965. Violent spore release in Helminthosporium turcicum in Nebraska. Phytopathology. 56:949-952.

MONTEITH, J. 1924. The leaf-spot disease of bluegrass. Bull. Green Sect. U.S. Golf Assoc. 4:172-173.

MORRISON, R. H. 1980. Leaf spot of Kentucky bluegrass in Minnesota: Misidentification of Drechslera poae as Drechslera dlctyoides. In: P. 0. Larsen and B. G. Joyner, eds. Advances in Turfgrass Pathology. Harvest Publications, Cleveland, Ohio. 149

MOWER, R. G. 1961. Histological studies of suscept-pathogen relations of Helminthosporium sativum, Helminthosporium vagans, and Curvularia lunata on leaves of 'Merlon* and common Kentucky bluegrass (Poa pra­ tensis L.). Ph.D. Thesis, Cornell Univ., N.Y. 150p.

NEELY, D. 1970. Persistance of foliar protective fungicides. Phyto­ pathology. 60:1583-1586.

NEELY, D., and E. B. HIMELICK. 1966. Simultaneous determination of fungistatic and fungicidal properties of chemicals. Phytopathology. 56:203-209.

PETERSON, L. J. 1959. Relations between inoculum density and infection of wheat uredospores of Puccinia graminis var. tritici. Phyto­ pathology. 49:607-614.

SLEESMAN, J., P. 0. LARSEN and J, SAFF0RD. 1974. Maintenance of stock cultures of Helminthosporium maydis (race T and 0). Plant Dis. Reptr. 58:334-336.

TORGENSON, D. C. 1969. Determination and measurement of fungitoxicity. Pages 93-123 in: D. C. Torgenson, ed. Fungicides an advanced treatise. Academic Press, London and New York. Vol. I. 697p.

VARGAS, J. M . , and R. D. WILCOXSON. 1969. Some effects of temperature and radiation on sporulation by Helminthosporium dictyoides on agar media. Phytopathology. 59:1706-1712.

WAGGONER, P. E. 1973. The removal of Helminthosporium maydis spores by wind. Phytopathology. 63:1252-1255.

WAGGONER, P. E., J. G. H0RSFALL, and R. J. LUKENS. 1972. Epimay: a simulation of southern corn leaf blight. Bull Conn. Expt. Sta. * New Haven. 84p.

WALLIN, J. R. 1973. Spore sampling of Helmithosporium maydis race T in 1971. Plant Dis. Reptr. 57:298-299.

WEIHING, J. L., S. G. JENSON, and R. I, HAMILTON. 1957. Helminthosporium sativum, a destructive pathogen of bluegrass. Phytopathology. 47:744-746.