CRITERIA FOR RANKING EVALUATIONS OF IR-4 ADVANCED …ir4.rutgers.edu/Biopesticides/Proposals/15A.pdf · 2. Experimental design, work plan and preliminary research. of 10 3. Does experimental

Reviewer’s Initials

CRITERIA FOR RANKING EVALUATIONS OF IR-4

ADVANCED STAGE BIOPESTICIDE PROPOSALS-2014 Proposal number/Title/PI: 15A, US Efficacy Trials for the Turf Bioherbicide, Sarritor, Hallett

The following criteria were established to assist the reviewers in selecting biopesticide projects for funding that: (1) are either in a more advanced stage of development (as opposed to exploratory or early stage of development) or involve expansion of the label; (2) have a high probability of being registered/marketed in a reasonable period of time; and (3) will be useful in meeting pest control needs involving minor crops (uses), including minor uses on major crops.

Criteria Score

(0 to 10 or 20)

1. Adequacy of investigators and facilities. of 10

2. Experimental design, work plan and preliminary research. of 10

3. Does experimental design allow to determine performance relative to conventional control practices and how the biopesticide might fit into IPM programs.

of 10

4. Evaluation of Budget of 10

5. Relevance of the proposal toward the development of data for

registration or label expansion of the biopesticide. of 10

6. Evidence of Efficacy. Positive supporting data provided. of 20

7. Probability of biopesticide being used by growers (factors such as effectiveness and economics of use rates should be considered). of 10

8. Other control measures currently available to control target pest. of 10

9. Probability of biopesticide being registered, time to registration,

and if label expansion, time to market. of 10 TOTAL* of 100

Funding Recommendation YES ____________ (Check appropriate line) NO ____________

MAYBE ____________ Note: Attach a comment page, should you have specific comments related to the proposal not covered in the above criteria. * There is a possibility of 10 points per criteria (except efficacy=20) for a total of 100 points. A rating of 0 means that the proposal does not meet the criteria at all, while a rating of 10 means it is ideal.

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IR-4 BIOPESTICIDE GRANTS COVER PAGE

2014

Proposal Number(For IR-4 Use): Principal Investigator: Steven G. Hallett Proposal Title: US Efficacy Trials for the Turf Bioherbicide Sarritor Institution: Purdue University Total dollars Requested $ 15,000.00

Enter each biopesticide /crop/ pest combination

No. Biopesticide and/or Conventional Product TRADE Name

Active Ingredient

Crop Pest (Weeds, Diseases, Insects)

1 Sarritor Sclerotinia minor IMI 344141

turfgrass Broadleaf weeds

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Proposal Title: US Efficacy Trials for Turf Bioherbicide Sarritor Address

Name Street City/State Zip Phone/Fax Email Address Project Director (Principal Investigator): Steve Hallett 915 West State Street West Lafayette 47907-2054 Ph. 765-494-7649

Fax. 765-494-0363 [email protected]

Administrative Contact: Carla Whiteman 615 West State Street West Lafayette 47907-2053 Ph. 765-494-6107 Fax. 765-496-1104

[email protected]

Financial Grant Officer: Carla Whiteman 615 West State Street West Lafayette 47907-2053 Ph. 765-494-6107 Fax. 765-496-1104

[email protected]

Authorized Grant Official: Amy Wright 625 Harrison Street West Lafayette 47907-2026 Ph. 765-494-8366 Fax. 765-496-1261

[email protected]

Individual Responsible for Invoicing: Carla Whiteman 615 West State Street West Lafayette 47907-2053 Ph. 765-494-6107 Fax. 765-496-1104

[email protected]

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I. Grant Stage - Advanced

Product is not currently registered with EPA, but a listing of all data and information submitted to EPA and EPA’s classification.pdf file is attached to this application.

II. Introduction (Limit 1 page) Include the objective, description of the pest problem and justification.

The monoculture turfgrass system provides a favorable environment for weeds. Weed species common to turfgrass vary with geographic regions, but many are common to more than one region. Weeds compete with turf for light, soil nutrients, soil moisture and physical space and can encroach turf. Seventy-three grass and grass-like and 145 broadleaf species are classified as weeds in turfgrass environments (McCarty et al. 2001). A combination of two or three herbicides is normally recommended to control a wide spectrum of broadleaf weeds. Repeated applications of dicamba or phenoxy herbicides such as 2,4-D and mecoprop or a 3-way product such as “Weed B Gon™” are extensively used for dandelion and other broadleaf control. As public concern increased around possible adverse health and environmental effects of lawn pesticides, bans or restrictions on their uses have increased. Even where amenity pesticide restrictions do not exist, interest in “natural” strategies for turfgrass management continues to grow. The search for biological control options has intensified.

SARRITOR bioherbicide was the first registered Microbial Pest Control Product (MPCP) in Canada for the control of Taraxacum officinale (dandelion) and other broadleaf weeds in turfgrass. Sclerotinia minor IMI 344141 is the active ingredient of SARRITOR. The fungus is cultured on ground barley and the bioherbicide granules are broadcast applied to weed infested turf. Favorable conditions for fungal germination and infection include 15-24C temperatures and >95% relative humidity. Disease develops quickly and complete kill of dandelion and other broadleaf weeds can be achieved within 7 days, about twice as fast as the standard auxinic herbicides. The product is compatible with normal lawn maintenance operations such as mowing, fertilization and irrigation. Registration of SARRITOR in the USA is pending. All toxicological and submitted documentation have been reviewed and accepted by EPA, but field trials in the United States are required to obtain data on SARRITOR that evaluates bioherbicide performance, environmental fate and persistence of S. minor IMI 344141 in representative turfgrass environments in the USA.

The IMI 344141 isolate was obtained from a lettuce field in Sherrington, Quebec but the life cycle, mode of action, moisture and temperature requirements, and host range of S. minor IMI 344141 were not different from S. minor “sensu lato” (Watson, 2007 ). However, persistence, survival and dissemination are much different when S. minor IMI 344141 is employed as an integrated biological control product. Isolate IMI 344141 can be phenotypically distinguished from the other tested S. minor (Shaheen et al., 2010) and a strain-specific molecular marker was developed to detect and monitor the S. minor IMI 344141 bioherbicide strain (Pan et al., 2010). When applied as a bioherbicide, S. minor (IMI 344141) did not persist into the following spring season in turf environments. This molecular detection method provides a mechanism to distinguish this isolate from related organisms and a tool to selectively monitor behavior of the biological control agent in the environment. When applied to turfgrass, S. minor IMI 344141 rarely produces sclerotia (melanized survival structures) and these sclerotia do not survive over

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winter. Mycelia of S. minor IMI 344141 that emerge from bioherbicide granules do not survive beyond 10 days in the turfgrass environment (Watson 2007).

III. Experimental Plan

1. The experiment will include the following treatments:

1. Untreated control

2. Conventional herbicide treatment (Weed B Gon Max at 1.7 kg ai/ha).

3. 1X bioherbicide 40 g/m2 (Sclerotinia minor (IMI 344141)

4. 2X bioherbicide -80 g/m2 (Sclerotinia minor (IMI 344141)

5. 1/2X herbicide (Weed B Gon Max 0.85 kg ai/ha) + 1X bioherbicide (40 g/m2 (Sclerotinia minor (IMI 344141)

6. Recent EPA registered biochemical pesticide (Fiesta Turf Weed Killer at 5.7 fl. oz./yd²)

Weed Control Products Being Evaluated:

1. SARRITOR (bioherbicide) (Sclerotinia minor (IMI 344141) 64.24%, other ingredients 35.76%, 300 infective units/g).

2. WEED B GON MAX (three way mix) (2,4-D (2,4-dichlorophenoxyacetic acid, 11.0%, 0.970 lb/gal), trichlopyr (3,5,6-trichloro-2-pyridinyloxyacetic acid, 1.12%, 0.097 lb/gal) and dicamba (3,6-dichloro-2-methoxybenzoic acid, 1.12%, 0.097 lb/gal).

3. FIESTA TURF WEED KILLER (Iron HEDTA (FeHEDTA) 26.52%, other ingredients 73.48%.

2. Crop and location: The experiment will be conducted on turf research fields at Purdue University, West Lafayette, IN, at Cornell University, Ithaca, NY, and at North Carolina State University, Raleigh, NC. Turf will be primarily Kentucky bluegrass (Poa pratensis) in West Lafayette, IN and Ithaca, NY and tall fescue (Festuca arundinacea) in Raleigh, NC.

3. Experimental design: Meter square plots of the 6 treatments will be replicated four times and arranged in a randomized complete block design.

4. Locations: The experiments will be conducted on turf research fields at Purdue University, West Lafayette, IN, at Cornell University, Ithaca, NY, and at North Carolina State University, Raleigh, NC.

5. Proposal fit into an IPM program. Recent bans on synthetic pesticides – including herbicides, insecticides, and fungicides – for cosmetic use in several Canadian provinces, have left lawn care professionals and residents with very few options to combat weeds in lawns, sports fields, parks, school grounds, and playgrounds. Sentiment in the United States may also lead to similar bans in the near

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future. For example, the city of Evanston, Ill., adopted the "Sustainable Pest Control and Pesticide Reduction Policy" in April 2010. It requires city employees, agents, and contractors to follow natural lawn care practices and prohibit the use of conventional pesticides. This adoption is a direct result of the municipality's desire to use more environmentally friendly products in areas where people may come into contact with the product. Several eastern states have taken a proactive stance with regards to potential regulation and have already taken measures to implement bans or restrictions in advance of any nationwide standards. Expanding community support for sustainable lawn practices will build and maintain demand for biopesticides in the United States.

Biopesticides include naturally occurring substances that control pests (biochemical pesticides), microorganisms that control pests (microbial pesticides), and pesticidal substances produced by plants containing added genetic material (plant-incorporated protectants). Fiesta is the only selective postemergence herbicide approved by the EPA's Biopesticide Division and labeled for broadleaf weed suppression in turfgrass. The active ingredient in Fiesta is iron chelate – FeHEDTA. It is quickly absorbed by the leaf tissues which turn black or brown. Contact-type action results in re-growth of many broadleaf weeds necessitating re-application to maintain control.

Sarritor bioherbicide (Sclerotinia minor IMI 344141) received full registration in Canada in 2010 for the control of dandelion and other broadleaf weeds in turfgrass. The first and only biological, 100% natural (nonchemical), compliant with all municipal by-laws, selective product for dandelion control was available for the commercial and domestic turfgrass markets. Sclerotinia minor IMI 344141 is an asporogenic plant pathogen that has biological control activity on dandelion and many other broadleaf weeds without damage to turfgrass species. The fungus is cultured on ground barley and the bioherbicide granules are broadcast applied to weed infested turf. Favorable conditions for germination and infection include 15-24C temperatures and >95% relative humidity. Disease develops quickly and complete kill of dandelion and other broadleaf weeds can be achieved within 7 days, about twice as fast as an industry-standard auxinic herbicide.

6. Data collection – Broadleaf weed control and turf health will be recorded at 7, 14, 21, 28 and 52 days after application (Abu-Dieyeh and Watson 2007). Symptoms of damage to dandelions and other broadleaf weeds will be visually estimated weekly for four weeks after application using a 0 to 10 visual scale compared to the control within the same mowing height and the same block, where 0 = not visually different from non-treated plots, 1 = 10-19% ….9 = 90-99% and 10 = 100% collapse of aboveground biomass. Data will be converted back to a percentage for analysis and presentation (after Franz et al. 1986 and Schnick et al. 2002). Turfgrass quality will be evaluated on a similar scale where 0 = turfgrass quality is indistinguishable from the non-treated areas, 1 = ~10% reduction is quality (discoloration, necrosis, or reduction in growth),… 9 = 90% of turfgrass damaged, 10 = 100% of the turf necrotic or dead.

At 7, 10, 21, and 28 days after application, the surface of bioherbicide and control plots will be search for the presence of sclerotia. Debris on the soil surface (1-2 cm depth) will be collected, air dried and out sourced for analyses. A strain specific molecular marker was developed to detect and monitor the Sclerotinia minor IMI 344141 bioherbicide strain (Pan et al. 2010). Soil samples (2 to 7 cm deep with soil

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sample core - 2cm diameter) are to be collected 5, 15, 30, 60, and 90 days after application in plots treated with the bioherbicide. Soil samples are also collected before application and 60 days after application from control plots in order to verify absence of IMI344141 in the field. These soil samples will also be out sourced for DNA extraction and PCR.

Data will be analyzed using ANOVA to determine if treatments differ from one another. Differences in treatment means will be determined using Tukey's test (SAS) at P = 0.05. Colby's (1967) test will be used to determine the nature of interactions among combination treatments.

7. Describe the pests to be controlled, the degree to which they are a problem in your state or region and the frequency that they occur (season long problem, every year, every few years).

Broadleaf weeds are common in most turfgrass sites every year and in every season. The most common and troublesome weeds in North Carolina turf include chickweed (Stellaria media), henbit (Lamium amplexicaule), clovers (Trifolium spp.), dandelion, cudweeds (Gnapthalium spp.) and violets (Viola spp.) (Webster 2008). Perennial broadleaf weeds such as dandelion, plantain (Plantago spp.) and white clover (Trifolium repens) are nearly ubiquitous in cool-season and transition zone lawns throughout North America. Other dominant species change with the seasons, with winter annuals (such as chickweed and henbit) dominating in the fall and spring; summer annual species (such as Chamaecyse and Oxalis) dominating in the warmer season. Test sites will be selected to contain representative weed species with emphasis on ubiquitous species such as dandelion and white clover.

8. Experiments will be conducted on naturally weed infested turfgrass. Where weed populations are inconsistent, weed populations will be augmented by disturbance and seeding or by transplanting.

9. What is the proposed start date and completion date? Also describe this in chronological order in the context of the experimental plan.

Site July – Aug 2014

Sept – Oct 2014

Nov – Dec 2014

Jan – Feb 2015

Mar – April 2015

May – June 2015

North Carolina

Prepare test sites

Treat Evaluate Sample

Evaluate Sample

Data recording and analysis

Final evaluations Data analysis

Final report

Indiana and New York

Prepare test sites

Treat Evaluate Sample

Evaluate Sample

Data recording and analysis

Final evaluations Data analysis

Final report

10. Test facilities where these studies will be conducted.

North Carolina experiments will be conducted on established turfgrass areas at the Horticulture Field Laboratory at North Carolina State University, Raleigh NC. The Horticulture Field Laboratory is a research experiment station at which Dr. Neal has maintained turfgrass areas and cultivated beds with established weed populations, specifically for use in such experiments. The site also has outdoor

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container research facilities, potting facilities and on-site composting (for plants and substrates used in container experiments). Access to the site when staff is not present, is limited to authorized personnel only. The nearest planting of a susceptible food crop would be about 100 meters from the test sites.

Indiana experiments will be conducted on established turfgrass areas at the William H. Daniel Turfgrass Research and Diagnostic Center, West Lafayette, IN. This research center has maintained turfgrass areas with established weeds such as dandelion, specifically for use in such experiments. Access to the site when staff is not present is limited to authorized personnel only. The nearest planting of a susceptible food crop is greater than 200 meters from the test sites.

At Cornell University, experiments will be conducted at the Bluegrass Lane Turf Research Facility in Ithaca, NY. The facility is comprised of several turf plots that simulate lawns, playing fields, putting greens, and sod. Three full time technicians maintain the research experiments at Bluegrass Lane and access to the site when staff is not present is limited to authorized personnel only. The location of the turf facility is located off the Cornell campus is not adjacent to any food crops. The nearest planting of a susceptible food crop would be 1 mile from the test site.

11. Budget:

Other Personnel: $3,663 An undergraduate student will serve on the project and salary support is requested at approximately $8/hour for 458 hours for the year. The undergraduate student will be supervised by Dr. Steve Hallet and will set up plots, apply and monitor bioherbicide performance and collect weed control efficacy data. Fringe Benefits: Undergraduate Student: 9.2% $337

Other Direct Cost:

Materials and Supplies: $500 The total request in supplies is based on our current research expenditures. Supplies required to do the work are culture media, plot management (stakes, etc.) and data collection). Travel: $500 Travel dollars are requested to travel to research plots from the main campus, and for partial support of one visit to meet with other researchers involved in the project at Cornell University.

Sub-Contracts: $10,000 Purdue will have a sub-contract with Cornell University and North Carolina State University. Please see the budget justifications for Cornell University and North Carolina State University below. Total Project Request of $15,000

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Cornell University Budget Justification Salary: Graduate Research Assistant (Kao-Kniffin): ($2,000) The funds will cover the summer salary of a graduate research assistant (GRA). The graduate assistant will be responsible for assisting with the inoculation and experimental set up, maintaining the plants, and assisting with data collection. Undergraduate hourly workers (DiTommaso): ($2,000) The funds will cover the salaries of undergraduate hourly workers that will assist the co-PIs and the graduate student will sampling, data collection, and sample analysis.

Other Direct Cost:

Land Rental: ($500) The requested funds will cover the cost of renting field plots at the Turfgrass Research Center.

Materials and Supplies: ($500): The requested funds cover the costs of supplies for the field study and sample analysis. Items include sampling bags, gloves, flags, and labware.

Total budget for Cornell is requested: $5,000

North Carolina State University Budget Justification

Salary: Partial salary (0.083 FTE) for a research technician who will be responsible to establishment, treatment, and maintenance of the research plots – including data recording, analysis and reporting. ($3,692) NCSU fringe benefit rate for staff is 30% of salary. ($1,108)

Other Direct Cost:

Materials and Supplies: Supplies related to establishing and maintaining experiments may include plot marking paint and stakes; mailing supplies (to send samples for analysis); personal protective equipment (such as chemical protective suits, nitrile gloves, rubber boots, eye protection, dusk masks); irrigation supplies; temporary nylon fencing (to restrict access to recently treated areas); maintenance chemicals (such as fertilizer, fungicides, insecticides); measuring devices (including: graduated cylinders, pipettes, pipette tips, weighing bags, balance); storage containers for inoculum, weed seeds, and related. ($200) Total budget for North Carolina State University is requested: $5,000

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12. Why product is needed.

Sarritor effectively suppresses top growth of dandelion, white clover, broadleaf plantain, bull thistle and other broadleaf weeds in turf. Based on the mode of action of Sclerotinia minor strain IMI 344141, the development of herbicide resistance is unlikely. The availability of Sarritor would enable further development of integrated and sustainable turf management practices, especially where the use of traditional chemical herbicides is not desirable. The alternative conventional and alternative biopesticide treatments are included in the proposed field experiments.

References

Abu-Dieyeh, M.H. and Watson, A.K. 2006 Effect of turfgrass mowing height on biocontrol of dandelion with Sclerotinia minor. Biocontrol Sci. and Technol. 16(5):509-524.

Abu-Dieyeh, M.H. and Watson, A.K. 2007. Population dynamics of broadleaf weeds in turfgrass as influenced by chemical and biological control methods. Weed Sci. 55(4):371-380.

Colby, S. R. 1967. Calculating synergistic and antagonistic responses of herbicide combinations. Weeds 15:20–22.

Frans, R. E, R. Talbert, D. Marx, and H. Crowley. 1986. Experimental design and techniques for measuring and analyzing plant responses to weed control practices. pp 29-46 In: Camper, D. ed. Research Methods in Weed Science, 3rd edition. Southern Weed Sci. Soc. Champaign, IL.

McCarty LB, Everest JW, Hall DW, Murphy TR. Yelverton F. 2001. Color atlas of turfgrass weeds. Chelsea, MI: Ann Arbour Press.269p

Pan, L., Ash, G.J., Ahn, B. and Watson, A.K. 2010. Development of strain specific molecular markers for the Sclerotinia minor bioherbicide strain IMI 344141, Biocontrol Sci. and Technol. 20(9), 939-959.

Schnick PJ, Stewart-Wade S, Boland G. 2002. 2,4-D and Sclerotinia minor to control common dandelion. Weed Science 50:173-178.

Watson AK (2007) Sclerotinia minor – biocontrol target or agent? Chapter 10 in Novel Biotechnologies for Biocontrol Agent Enhancement and Management (eds. M. Vurro and J. Gressel), Springer,205-211.

Watson, A.K. and Bailey, K. (2013) Taraxacum officinale Weber, Dandelion (Asteraceae). Ch. 58 in Biological Control Programmes in Canada 2001-2012, (eds. Mason P and Gillespie D), CABI Publishing, Wallingford, UK, pp. 383-391.

Webster, T.M. ed. 2008. The Southern States 10 most common and troublesome weeds in turf. (Table 7). P. 239. IN: Southern Weed Sci Soc. Proceedings 61:239. http://www.swss.ws/publications/proceedings/

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Page 1 of 3

SARRITOR®

[Alternate Brand Names:]

Granular Biological Herbicide

Active Ingredient: Sclerotinia minor IMI 344141*…………..………………………………..64.24 %

Other Ingredients : .……………………………………………….……..………………..…..35.76 %

Total : ……………………………………………………………..……………..………...…..100.0 %

* Contains a minimum of 300 infective units/gram

KEEP OUT OF REACH OF CHILDREN

CAUTION

FIRST AID

If swallowed:

Call a poison control center of doctor immediately for treatment advice.

Have person sip a glass of water if able to swallow.

Do not induce vomiting unless told to do so by the poison control center or doctor.

Do not give anything by mouth to an unconscious person.

If on skin or clothing:

Take off contaminated clothing.

Rinse skin immediately with plenty of water for 15-20 minutes.

Call a poison control center or doctor for treatment advice.

If inhaled:

Move person to fresh air.

If person is not breathing, call 911 or an ambulance, then give artificial respiration, preferably

mouth-to-mouth, if possible.

Call a poison control center or doctor for further treatment advice.

If in eyes:

Hold eye open and rinse slowly and gently with water for 15-20 minutes. Remove contact

lenses, if present, after the first 5 minutes, then continue rinsing.

Call a poison control center or doctor for treatment advice.

Hot Line Number Have the product container or label with you when calling a poison control center or doctor, or going for treatment.

For emergency information, call a local poison control center at 1-800-222-1222.

See back panel for additional precautionary statements

EPA Reg. No. XXXXX-X • EPA Est. No. XXXXX-PA-1

Manufactured by: Distributed by: 4260864 Canada Inc. W.F. Stoneman Company

104 Rhapsodie, Notre-Dame-de-l’Ile-Perrot, Post Office Box 465 QC J7V 8P1 Canada McFarland, WI 53558-0465

e-mail: [email protected]

Net Contents: 15 pounds

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mailto:[email protected]

mailto:[email protected]

Page 2 of 3

PRECAUTIONARY STATEMENTS

HAZARDS TO HUMANS AND DOMESTIC ANIMALS: CAUTION. Causes moderate eye irritation. Harmful if swallowed, or absorbed through skin. Avoid contact with skin, eyes or clothing. Wear protective eyewear. Wash thoroughly with soap and water after handling and before eating, drinking, chewing gum, using tobacco or using the toilet. Remove and wash contaminated clothing before reuse.

PERSONAL PROTECTIVE EQUIPMENT (PPE): Applicators and other handlers must wear: • Long-sleeved shirt and long pants. • Chemical resistant gloves. • Shoes plus socks. • Protective eyewear.

Mixers/loaders and applicators must wear a NIOSH-approved respirator with any N, P, R, or HE filter. Repeated exposure to high concentrations of microbial proteins can cause allergic sensitization. Follow manufacturer’s instructions for cleaning/maintaining PPE. If no such instructions for washables, use detergent and hot water. Keep and wash PPE separately from other laundry. ENGINEERING CONTROLS: When handlers use closed systems, enclosed cabs, or aircraft in a manner that meets the

requirements listed in the Worker Protection Standard (WPS) for agricultural pesticides [40 CFR 170.240(d)(4-6)], the handler PPE requirements may be reduced or modified as specified in the WPS. IMPORTANT: When reduced PPE is worn because a closed system is being used, handlers must be provided all PPE specified above for “applicators and other handlers” and have such PPE immediately available for use in an emergency, such as a spill or equipment break-down.

USER SAFETY RECOMMENDATIONS

Users should

Remove clothing/PPE immediately if pesticide gets inside. Then wash thoroughly and put on clean clothing.

Remove PPE immediately after handling this product. Wash the outside of gloves before removing. As soon as possible,

wash thoroughly and change into clean clothing.

ENVIRONMENTAL HAZARDS: For terrestrial uses: Do not apply directly to water, or to areas where surface water is present or to intertidal areas below the mean high water mark. Do not contaminate water when cleaning equipment or disposing of equipment wash waters or rinsate.

DIRECTIONS FOR USE

It is a violation of Federal law to use this product in a manner inconsistent with its labeling. Do not apply this product in a way that will contact workers or other persons, either directly or through drift. Only protected handlers may be in the area during application. For any requirements specific to your State or Tribe, consult the State or Tribal agency responsible for pesticide regulation.

Read the entire label before using.

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Page 3 of 3

NON-AGRICULTURAL USE REQUIREMENTS

The requirements in this box apply to uses of this product that are NOT within the scope of the Worker Protection Standard for agricultural pesticides (40 CFR Part 170). The WPS applies when this product is used to produce agricultural plants on farms, forests, nurseries or greenhouses. Keep unprotected persons out of treated areas until sprays have dried.

GENERAL INFORMATION: For use against broad leaf weeds in turf on home lawns, commercial lawns, golf courses, municipal parks, and sod farms. WEEDS CONTROLLED: Dandelion (Taraxacum officinale), plantain, (Plantago major), clover (Trifolium repens), thistle (Cirsium spp. ), and other broadleaf weeds.

APPLICATION DIRECTIONS: Apply broadcast with a drop spreader onto surface of actively growing weed infested turf at a rate of 9 to 13 pounds per 1000 square feet or spot apply directly on individual weeds at a rate of ½ to 1 teaspoon per plant. During hot dry weather, apply in the evening and irrigate thoroughly after 12 hours. Avoid contact with desirable broadleaf garden plants and scrubs, direct contact may damage non-target plants. A 3 day post-treatment no-mowing period is recommended.

STORAGE AND DISPOSAL Do not contaminate water, food or feed by storage and disposal.

PRODUCT STORAGE

Store in a dry, cool place out of direct sunlight and away from heat sources at 35 to 40 °F. Product is stable for up to

8 months when stored frozen. After opening, if product some product is remaining, reseal the original packaging

and use within one month.

PRODUCT DISPOSAL

To avoid wastes, use all material in this container by application according to label directions. If wastes cannot be

avoided, offer remaining product to a waste disposal facility or pesticide disposal program (often such programs are

run by state or local governments or by industry).

CONTAINER DISPOSAL Nonrefillable container. Do not reuse or refill this container. Completely empty container into application equipment

by shaking and tapping sides and bottom to loose clinging particles. Then offer for recycling if available or dispose

of empty container in a sanitary landfill or by incineration. Do not burn, unless allowed by state and local

ordinances.

WARRANTY - 4260864 Canada Inc. warrants that the product conforms to the description on the label and is

reasonably fit for the purposes set forth on the label, when used according to directions under normal use conditions.

Neither this warranty nor any other warranty of merchantability or fitness for a particular purpose, expressed or

implied, extends to the use of this product contrary to the label instructions; the buyer assumes the risk of any such

uses.

Batch No.

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http://en.wikipedia.org/wiki/Taraxacum_officinale

http://www.msuturfweeds.net/details/_/white_clover_28/

http://www.msuturfweeds.net/details/_/canada_thistle_18/

Effect of turfgrass mowing height on biocontrol ofdandelion with Sclerotinia minor

MOHAMMED H. ABU-DIEYEH & ALAN K. WATSON

Department of Plant Science, McGill University, Ste-Anne-de-Bellevue, Quebec, Canada

(Received 18 August 2005; returned 28 September 2005; accepted 8 November 2005)

AbstractThe fungus, Sclerotinia minor Jagger is under development as a bioherbicide for control ofdandelion and many broadleaf weeds in turfgrass environments. The effect of S. minor ondandelion survival was evaluated under different mowing heights and compared with acommonly used herbicide KillexTM. In the greenhouse, the onset of symptoms was more rapid,foliar damage was more severe, and the reduction of aboveground biomass and root biomasswas greater for the bioherbicide than the herbicide. The bioherbicide reduced root biomass]/10-fold compared with untreated plants. Under high weed infestation levels in the field,S. minor caused a greater initial reduction of dandelion density than did the herbicide during the2-week post-application period, although reductions were greater in herbicide treated plots by6 weeks after application. Over the growing season, S. minor and the herbicide had similarsuppressive effects on dandelion density except under the closest mowing height (3�5 cm).After treatment, close mowing favored dandelion seedling recruitment and the biocontrol hadno residual activity. Survival of dandelion roots was significantly less after spring than falltreatment of S. minor and season long mowing at the close height significantly reduce rootsurvival. Close mowing may be detrimental for S. minor applications on heavily infesteddomestic lawns and amenity grassland areas.

Keywords: Bioherbicide, biological weed control, clipping, fungus, mowing, Sclerotinia minor,

Taraxacum officinale, turfgrass, weed control

Introduction

Broadleaf weeds reduce the quality and quantity of turfgrass and disrupt its

visual uniformity (McCarty et al. 2001). Dandelion, Taraxacum officinale Weber, is

a strong competitive perennial weed that infests turfgrass environments and is

common in home lawns, turfgrass swards, pastures, forages, golf courses, athletic

fields, wooded areas, and roadside verges (Stewart-Wade et al. 2002a). It is an

undesirable plant, causing aesthetic problems during flowering and seed production.

In 1992, a survey of turfgrass specialists estimated that there were 7.3 million hectares

of lawn in the United States (Elmore 1994). The lawn care industry in North America

Correspondence: Alan Watson, Department of Plant Science, McGill University, 21,111 Lakeshore Road,

Ste-Anne-de-Bellevue, Quebec, Canada H9X 3V9. Tel: 1 514 398 7858. Fax: 1 514 398 7897. E-mail:

[email protected]

First published online 14 March 2006

ISSN 0958-3157 print/ISSN 1360-0478 online # 2006 Taylor & Francis

DOI: 10.1080/09583150500532725

Biocontrol Science and Technology, 2006; 16(5): 509�524 15A

has been expanding at a rate of 5�8% per year (United States Environmental

Protection Agency 1999). The need for regular management and maintenance of

these areas creates a massive turfgrass industry in North America and Europe. In

2001, 78 million households in the US used home and garden pesticides (United

States Environmental Protection Agency 2004). Herbicides accounted for the highest

usage of pesticides in the home and garden sector with users spending 632 million

dollars for over 36 million kg applied on lawns and gardens. 2,4-D (2,4-dichloro-

phenoxy acetic acid) was the most widely used pesticide in the home and garden

sector.

Repeated applications of phenoxy herbicides such as 2,4-D and mecoprop ((9/)-2-

(4-chloro-2-methylphenoxy) propanoic acid), dicamba (3,6-dichloro-2-methoxyben-

zoic acid), or combination products such as KillexTM have been widely used for

dandelion control (Anonymous 2005). Environmental and public health concerns

about pesticides, especially the use of chemical herbicides in turfgrass for aesthetics,

have led to the banning or severe restrictions on the use of pesticides in many

regions of Canada (Cisar 2004). Consequently, alternatives to chemical herbicides

are being sought (Neumann and Boland 2002; Stewart-Wade et al. 2002b; Zhou et al.

2004).

Sclerotinia minor Jagger is an asporogenic ascomycete plant pathogen that has

biocontrol potential for dandelion in turfgrass, and several formulations of S. minor

have been shown to have biological control activity (Ciotola et al. 1991; Riddle et al.

1991; Briere et al. 1992; Stewart-Wade et al. 2002b; Abu-Dieyeh & Watson 2005).

S. minor mycelium in sodium alginate granules (Briere et al. 1992) and mycelial-

colonized barley grits (Stewart-Wade et al. 2002b) effectively controlled dandelion

and broadleaf plantain (Plantago major L.) without damage to turfgrass species. When

applied to turfgrass, S. minor IMI 344141 rarely produces sclerotia (melanized

survival structures), and these sclerotia do not survive over winter. Mycelia of S. minor

IMI 344141 does not survive beyond 10 days in the turfgrass environment. Field and

greenhouse studies confirmed that turfgrass species are not susceptible to S. minor

IMI 344141. Independent toxicological studies have established that S. minor IMI

344141 is neither toxic nor pathogenic to humans, birds, fish, daphnia, honey bees,

earthworms, or wild animals.

Extension recommendations for turfgrass often indicate that a dense, healthy

turfgrass stand is the best defense against weed colonization and can be managed

by proper mowing, watering, and fertilization (McCarty et al. 2001; Busey 2003).

A sensible recommendation but the methods are often stated in generalities and are

not based on scientific research (as stated by Busey 2003). Mowing is a major cultural

practice for turf maintenance. Lower mowing heights can cause additional stress for

certain grass species and/or broadleaf weeds, especially in summer (as reviewed by

Busey 2003; Hatcher & Melander 2003), and higher canopies may maintain higher

humidity levels close to the soil surface for longer time periods, which can be

advantageous for pathogen activity (Gielser et al. 2000). When a fungal pathogen was

combined with simulated mowing, weed suppression was greater than the fungus

alone (Green et al. 1998; Kluth et al. 2003). In this research, the effect of turfgrass

mowing height on the efficacy of a granular formulation of S. minor to suppress

dandelion in a suburban lawn environment was evaluated.

510 M. H. Abu-Dieyeh & A. K. Watson 15A

Materials and methods

Production and formulation of S. minor

S. minor (IMI 1344141) was isolated from diseased lettuce plants (Lactuca sativa L.)

from southwestern Quebec in 1988, and stock cultures maintained as sclerotia at 48C.

When required, sclerotia from a stock culture were washed twice in sterile distilled

water, placed in 70% ethanol for 40 s, transferred to 1% hypochlorite solution for

3 min, rinsed twice with sterile distilled water, and set to dry on sterilized filter paper.

The surface sterilized sclerotia were transferred aseptically onto potato dextrose agar

(PDA, DIFCO Laboratories, Detroit, MI) plates and incubated for 4�5 days at 209/

18C. Five agar plugs (5 mm diameter), from the actively growing margin of colonies

on PDA were transferred to 100 mL of a modified Richard’s solution (MRS) having

the following constituents L�1: 10 g of sucrose, 10 g of KNO3, 5.0 g of KH2PO4,

2.5 g of MgSO4 �/7H2O, 0.02 g FeCl3 �/6H2O, and 150 mL V-8 juice (Campbell Soup

Company Inc.) in 250-mL Erlenmeyer flasks. Cultures were incubated for 5 days on a

rotary shaker at 60 rpm at room temperature (209/18C). The grown mycelium were

collected into a sterilized blender cup (Waring Commercial, Torrington, CT) and

homogenized gently with two 20-s bursts and then inoculated onto autoclaved barley

(Hordeum vulgare L.) grits. For this, whole barley grains were ground and sieved to

1.4�2.0 mm diameter grits. Three hundred g of barley grits were transferred into

autoclavable bags with a breathable patch 44�/20.5 cm, 0.02 mm filter: 24 mm

(SunBag, transparent, Sigma-Aldrich, Montreal, QC). Distilled water (210 mL) was

placed into each of the bags and autoclaved at 1218C for 20 min. After autoclaving,

the bags were allowed to cool and a 15-mL of the liquid S. minor mycelial culture was

transferred aseptically into each bag. Inoculated bags were incubated at 209/18C in

the dark and shaken on the third to sixth days of incubation. The contents of each bag

were then dried separately by spreading the colonized barley grits onto mesh trays for

12 h under a laminar flow. The dried inocula (aw 0.4) were placed in plastic bags

(PolyBags, 17.5�/40�/7.5 cm, Gerrity Corrugated Paper Products, Concord, ON)

and the bags were sealed, and stored at 48C. These S. minor granular formulations

were used in our experiments after 2�4 weeks of storage. The viability of the S. minor

preparations was verified by incubating 10 granules from each bag on PDA plates.

Colony diameters were measured after 24 and 48 h of incubation at 209/18C in the

dark. Additionally, 10 granules from each bag were placed onto excised dandelion

leaves maintained on moist sterile filter papers in Petri dishes and incubated at 208C in

the dark; one granule/leaf. The diameter of the lesions caused by the fungus was

measured after 24 and 48 h of incubation. Previous unpublished quality control

studies indicated viable batches to have colony diameter of 14�30 mm after 24 h,

and 40�70 mm after 48 h and virulent batches to have an average lesion diameter

�/15 mm after 48 h of incubation. S. minor preparations were only used in our

experiments if they met these criteria.

Effect of S. minor and mowing height on dandelion control in the greenhouse

Dandelion seeds collected in spring 2002 from lawns on the Macdonald campus,

McGill University, Ste-Anne-de-Bellevue, QC and stored at 48C were sown onto

potting soil (2/3 black pasteurized soil and 1/3 Pro-mix (Premier Promix, Premier

Horticulture Ltee, Riviere-du-Loup, QC) in plastic containers (40�/32�/20 cm,

Mowing effect on performance of a turf bioherbicide 51115A

L�/W�/D, 19 L capacity; Sterilite Inc., Montreal, QC). One week after germination,

seedlings were thinned to four equidistant seedlings per container. Two weeks

after germination of the dandelion seeds, 2.5 g of a commercial grass seed mixture

[30% Kentucky bluegrass (Poa pratensis), 40% creeping red fescue (Festuca rubra L.

var. rubra) and 30% turf type perennial ryegrass (Lolium perenne L.), C.I.L.†

GolfgreenTM, Brantford, ON] was scattered over the surface of each container.

Grass was cut weekly with hedge shears (PlantSmart, Wal*Mart, Canada),

commencing 3 weeks after grass sowing to a height of 5, 10, 15, or 20 cm. Four

weed control treatments were imposed: (1) untreated, (2) a spot application of

0.2 g plant�1 non-colonized autoclaved barley grits, (3) a spot application of

0.2 g plant�1 barley granular formulation of S. minor , and (4) a broadcast foliar

application of KillexTM at 1.7 kg a.i ha�1 (25 mL of the 0.6% original concentration

per container). The herbicide was applied with a 1.18-L vacuum sprayer (Home and

Garden sprayer. Model no 1998. RLF10-Master Premium. Root-Lowell Manufactur-

ing Co., Lowell, MI). The weed control treatments were applied 3 weeks after

initiating of grass cutting heights and thus dandelion plants were 8 weeks old. Prior to

weed control treatment applications, all plants were misted lightly with water to aid

barley grit adhesion with dandelion leaves.

The plants were grown in the greenhouse at 209/28C with 15 h of light/day at a

minimum photon flux density of 3509/50 mmoL m�2 s�1. Plant containers received

programmed drip irrigation of 150 mL 3�/day�1. To enhance the establishment of

the plants, a 15:15:30 N�P2O5�K2O vegetable fertilizer with micronutrients

(Plantex†, Plant Product Co, Brampton, ON) was applied at 3.5 g L�1 when the

dandelions were 5 weeks old.

Symptoms of damage to dandelions were visually estimated weekly for 4 weeks after

application using a 0�10 visual scale compared to the control within the same mowing

height and the same block, where 0�/B/9%, 1�/10�19% . . . 9�/90�99% and 10�/

100% collapse of aboveground biomass. Data were converted back to a percentage for

analysis and presentation. Plant regrowth was measured as a reduction in percent

damage by estimating the biomass of new leaves produced post-inoculation compared

to the control within the same mowing height and the same block. Damage estimates

of the four plants in each container were averaged and analyzed as one experimental

measure. The number of post inoculation dandelion plants that survived was recorded

weekly for 6 weeks. Six weeks after application, all of the dandelion plants were

carefully removed from the soil to extract their entire tap root. The roots were

thoroughly washed and dissected above the crown, separating above ground and

below ground biomass. All leaf or root biomass from each container was bulked,

placed in paper bags, oven-dried at 808C for 72 h, and then weighed.

The experiment was a split-plot design with five replicates and was conducted twice

through time (January 2003 and January 2004). Main plots were weed control

treatments and subplots were grass heights. In the repeat trial in 2004, the non-

colonized autoclaved barley grits treatment was omitted and excluded from the pooled

analysis as no significant differences were obtained for any of the studied parameters

when compared with the untreated treatment in the first trial. Data for each parameter

from the two experimental trials were subjected to the Bartlett test of SAS (SAS

Institute Inc., Cary, NC, 2002) to test for homogeneity of variances. Data for all

parameters were homogeneous, thus they were pooled. The main effects of weed

control treatments, mowing heights, and their interaction were determined using


ANOVA of SAS, and the effects of time on aboveground damage data were analyzed

using SAS GLM procedure of repeated measures. The means were separated using

the Tukey test at P�/0.05 (SAS Institute Inc., Cary, NC, 2002).

Effect of S. minor and mowing height on dandelion control in the field

Two field experiments were conducted, one in each of two different sites on the

Macdonald Campus of McGill University, in Ste-Anne-de Bellevue, QC (45825?Nlatitude, 73855?W longitude, 39.00 m elevation). The climate data for the 2 years of

study (2003 and 2004) are summarized in Table I. Study Site-1 (2003 experiment)

was approximately 600 m2 on a loamy sand soil (coarse sand�/9%, fine sand 73%,

silt�/12%, clay�/6%) with a pH of 7.15 and 8% organic matter. The lawn was

established in 1973 and received low maintenance management throughout its history

except for repeated mowing during the growing season (May to October). The grass

sward was approximately 60% Kentucky blue grass, 30% perennial ryegrass, patches

of timothy (Phleum pratense L.), and rare occurrences of annual blue grass (Poa annua

L.). The lawn flora was highly diversified with broadleaf weeds (17 species were

observed throughout the study period), and the dominant weed species was dandelion

(Taraxacum officinale) with 50�60 plants m�2 recorded prior to the spring treatment

application.

Study Site-2 (2004 experiment) was located in an open lawn area of approximately

600 m2 with low human disturbance, established in 1980. The turf at Site-2 was in

better visual quality than in Site-1 with mainly Kentucky blue grass (�/90%) and 10%

red fescue (Festuca rubra L.). Herbicides had not been applied for the past 10 years

and the major broadleaf species were dandelion (60�70 plants m�2), white clover

(Trifolium repens L.) and broadleaf plantain (Plantago major L.). The soil was loamy

Table I. Weather data for Ste-Anne-de-Bellevue, Quebec during the 2 years of study 2003 and 2004.

Environment Canada Meteorological Data. Ste-Anne-de-Bellevue Station.

Temperature (8C) RH% Rainfall (mm)

Month/year Average Min Max Average Min Max Total monthly

May/03 13 2 28 70 15 100 136

May/04 7 �/1 29 68 22 100 123

Jun/03 17 7 32 70 27 100 124

Jun/04 17 4 29 65 26 99 226

Jul/03 20 11 30 73 27 100 357

Jul/04 20 11 31 75 30 99 574

Aug/03 22 14 29 79 35 100 53

Aug/04 19 8 29 78 45 100 182

Sep/03 17 5 29 77 31 100 416

Sep/04 16 5 26 79 41 100 96

Oct/03 7 �/2 29 80 31 100 222

Oct/04 9 2 25 77 33 100 85


sand (12% coarse sand, 75% fine sand, 7% silt and 6% clay) with a pH of 7.2 and

6.6% organic matter.

Both field experiments were established in May and maintained until the end of

October. Study sites (20�/30 m) were marked with metal posts and plastic ropes and

the corners of each plot were permanently marked by wooden sticks to maintain plot

integrity for the duration of the study. The area of the experimental unit (plot) was

1 m2 with 0.8 m alleys between plots. The distance between any two blocks was

2�3 m. The experimental design was a split plot with four replicates. Mowing heights

were the main plots and weed control treatments were the subplots. The three levels of

mowing heights, 3�5, 7�10, and 12�15 cm were initiated 2 weeks prior to weed

control treatment applications. Plots were mowed weekly, except during the 2-weeks-

post-weed control treatment period, with a gas powered rotary push mower. Grass

clippings were returned during July and August to act as a source of nitrogen (Kopp &

Guillard 2002), but removed during other months in the 6-weeks post-treatment

periods to avoid cross contamination between blocks.

Four levels of weed control were imposed on 15 May and on 15 September at

each study site: (1) untreated control, (2) a broadcast foliar application of KillexTM at

1.7 kg a.i. ha�1 (200 mL m�2 of 0.6% of original concentration), (3) a broadcast

application of 60 g m�2 granular formulation of S. minor , and (4) a broadcast

application of 120 g m�2 granular formulation of S. minor . The herbicide was applied

onto the grass surface using a 1.18-L vacuum sprayer. The S. minor formulation was

applied using a 200-mL plastic bottle fitted with a perforated lid (�/10 mm diameter)

with suitable openings to pass the barley grits. If there was no rainfall on the day of

application or the grass was not wet, the entire field was sprinkler irrigated for 2 h

prior to late afternoon treatment applications. No additional irrigation or fertilization

management was applied during the course of the study.

The number of dandelions was counted in each plot the day before weed control

treatments, and on a single day in the last week of each month thereafter. In order to

monitor post treatment recovery of dandelions, 10 dandelion plants in each of the

fungal treated plots were randomly marked using white colored pins prior to treatment

applications. In Site-2, the number of dandelion seedling recruits versus mature plants

in each plot was counted every 2 weeks starting 2 weeks after application (30 May)

and continuing to the end of June.

Dandelion density data were adjusted using the ‘Before�After, Control-Impact

(BACI) equation’ (Green 1979) to overcome spatial heterogeneity data differences

from pretreated time (15 May) and also to overcome the effect of progression in time.

BACI value� (At=Bt)=(Ac=Bc)�100 (1)

where B is the dandelion density at the time of treatment (before the impact), A is the

dandelion density after treatment (after the impact), t is the treatment, and c is the

control. The control used for comparison was the untreated plot at the same mowing

height in the same block. To realize the effect of mowing heights on dandelion in

untreated plots, the value obtained from the three mowing heights within the same

block were averaged and used as control in equation 1.

Data for each time period from the 2 years (two study sites) were subjected to

Levene’s procedure (SAS Institute Inc., Cary, NC, 2002) for testing homogeneity of

variances using three variances (the three levels of mowing heights). Because of

differences in environmental conditions between years and/or locations, monthly data


were heterogeneous except for the spring pre-treatment application (mid-May);

2 weeks post-application (end of May); 6 weeks post-application (end of June), and for

post-control season average. Therefore, the season average, the 2 and 6 weeks post-

application data from the two experiments were pooled and treated as one experiment.

The 120-g m�2 S. minor treatment was excluded from the analysis as no significant

differences were obtained in comparison with the 60-g m�2 treatment for all studied

parameters. Normality for each parameter was tested on model residuals using the

Shapiro�Wilk test (SAS Institute Inc., Cary, NC, 2002). Data were analyzed using

ANOVA to determine the significant interactions among mowing heights and different

weed control treatments. Differences in treatment means were determined using

Tukey’s test (SAS) at P�/0.05.

Results

Effect of S. minor and mowing height on dandelion control in the greenhouse

Eight-week-old dandelions (six to eight leaves) were highly susceptible to the S. minor

application. The above ground biomass in all treated plants collapsed during the first

week after application (Figure 1). Symptom expression was more rapid with S. minor

and at 1-month-post application it was more effective than the herbicide (KillexTM).

The herbicide and the S. minor treatments caused highly significant (P 5/0.01) above

Sclerotinia minor (a)

Killex(b)

Untreated

Weeks post application

0 1 2 3 4

% A

bo

ve g

rou

nd

dam

age

of

dan

del

ion

0

20

40

60

80

100

120

5 cm mowing height10 cm15 cm20 cm

Figure 1. Effect of mowing heights and weed control treatments on above ground damage (%) of dandelion

in a grass planting. Within each post-application time, the means of the three weed control treatments are

significantly different at the 1% level according to Tukey test. (a) Spot application of a granular formulation

of S. minor at 0.2 g plant�1. (b) Broadcast foliar application of KillexTM herbicide (2,4-D, mecoprop and

dicamba) at 1.7 kg a.i. ha�1.


ground biomass damage to the dandelion plants. S. minor caused significantly more

damage than the herbicide over the entire study period (Figure 1).

Mowing height had no significant effect on dandelion above ground damage in any

of the weed control treatments (Figure 1), but the above ground and root biomass of

untreated dandelion were significantly reduced under the 5�10 cm compared with

15�20 cm mowing heights (Figure 2A,B). At 6 weeks post-application, both S. minor

and the herbicide caused highly significant (P 5/0.01) above ground and root biomass

reduction compared with untreated plants. Biomass reductions were greater for

S. minor (Figure 2A,B). The plants surviving the initial S. minor treatment had short,

weak roots and few leaves sprouted from the crown (Figure 3). Regrowth following

S. minor treatments was least when combined with the closest mowing height (5 cm)

(Figure 4).

Effect of S. minor and mowing height on dandelion control in the field

The greatest effect of S. minor was observed 2 weeks after application when dandelion

densities declined by 65�95%. Under the 7�10 and 12�15 cm mowing heights, the

effect of the S. minor (60 g m�2) was significantly greater than that of the herbicide,

but at the 3�5-cm mowing height, the effect of S. minor and the herbicide were similar

2 weeks after application (Figure 5A).

Six weeks after application, dandelion densities within untreated plots were reduced

by mowing at 3�5 cm compared to mowing at 7�10 or 12�15 cm (Figure 5B).

Dandelion population densities were reduced more by the herbicide than by S. minor

at all mowing heights 6 weeks after application (Figure 5B). Under close mowing

(3�5 cm), dandelion densities in S. minor-treated plots increased from 35 to 80%

between 2 and 6 weeks post-application (Figure 5B).

5 10 15 20

Ab

ove

gro

un

d d

ry m

atte

r (g

)

0.0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

Mowing height (cm)

(A) (B)

Untreated

Killex(b)

Sclerotinia minor (a)

aa

ab

bc

bcdbcd

cde

de

e e e e

5 10 15 20

Ro

ot

dry

mat

ter

(g)

0.0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

a

a

b

b

b

b

b

b

cccc

Figure 2. Effect of mowing heights and weed control treatments on aboveground (A) and root (B) biomass

of dandelion, 6 weeks after treatment application. Mowing heights were initiated 1 month prior to treatment

application and maintained throughout the study period. Within the same graph, bars with similar letters

are not significantly different at the 5% level according to Tukey test. (a) Spot application of a granular

formulation of S. minor at 0.2 g plant�1. (b) Broadcast foliar application of KillexTM herbicide, at

1.7 kg a.i. ha�1.


The BACI post application average of dandelion population densities over the

entire growing season (Figure 5C) illustrates the recovery ability of the dandelion

population and also the accumulated effect of mowing. While the effect of mowing

heights was concealed under the strong effect of the chemical herbicide treatment, it

aa

ab

b

a

a a

Mowing height (cm)

5 10 15 20

% o

f reg

row

th fr

om r

oots

0

10

20

30

40

50

60

70

b

6 weeks post inoculation

8 weeks post inoculation

Figure 4. Effect of mowing height on dandelion root regrowth, 6 and 8 weeks after S. minor inoculation.

Within each post-application time, bars with similar letters are not significantly different at the 5% level

according to Tukey test.

Figure 3. A comparison of the above and below ground biomass between untreated dandelion (two plants

above) and dandelion regrown 6 weeks after inoculation with S. minor (three plants below).


was apparently able to interact significantly with seasonal average dandelion density in

untreated and fungal treated plots (Figure 5C).

Prior to the spring treatment, approximately 50% of the dandelion plants in all plots

were new seedling recruits (Figure 6). In untreated plots, these recruits decreased

gradually to approximately 10�20% of the population by 6 weeks after application

(Figure 6) mostly due to rise in temperature (Table I). In herbicide treated plots, no

seedlings were observed after 2 and 4 weeks post-application and very few at 6 weeks

after application. In S. minor treated plots, the ratio of recruitments to mature plants

was highly variable according to time and mowing factors. Within the first 2 weeks

post-application, no seedlings were observed under the 7�10 and 12�15 cm mowing

heights but a small number (less than 5 m�2) was reported under the closest mowing

height (Figure 6). After 4 weeks, the large recruitment of seedlings under the closest

mowing height resulted in significantly more mature plants after 6 weeks compared

with the other mowing heights (Figure 6).

3-5 cm 7-10 cm 12-15 cm

Dan

del

ion

den

sity

(B

AC

I (3)

valu

es)

Dan

del

ion

den

sity

(B

AC

I (3)va

lues

)

0

20

40

60

80

100

120

140

3-5 cm 7-10 cm 12-15 cm0

20

40

60

80

100

120

140

aa

bb

cc

d d d

a

ababc

bc

bcd

d

e e

ab

A

3-5 cm 7-10 cm 12-15 cm0

20

40

60

80

100

120

140

b

d

c

a

dd

a

d d

Untreated

Sclerotinia minor (2)Killex(1)

B

Mowing heights

C

Figure 5. Effect of mowing heights and weed control treatments on post application dandelion density after

2 weeks (A), 6 weeks (B), and season average (C). Mowing heights were initiated 2 weeks prior to spring

application and maintained throughout the experiment. Within the same graph, bars with similar letters are

not significantly different at the 5% level according to Tukey test. (1) Broadcast foliar application of KillexTM

herbicide, at 1.7 kg a.i. ha�1. (2) Broadcast of a granular formulation of S. minor at 60 g m�2. (3) Before�After, control-Impact�/(At /Bt )/(Ac /Bc )�/100, B is the density before treatment; A is the density after

treatment; t is the treated; and c is the control.


SeedlingsMature plants

2-weeks post application

0

10

20

30

40

50

60

70

AB

a

d

B

cC

A

a

AB

d d

C

b

AB

d

AB

dC


0

10

20

30

40

50

60

70

b

c

a a

c

b

a

c

b

A

E

B

A

DE

BC

A

CDBCD

3-5 cm

Untreated Herbicide S. minor Untreated Herbicide S. minor

7-10 cm

Untreated Herbicide S. minor

12-15 cm

3-5 cm


7-10 cm


12-15 cm

3-5 cm


7-10 cm


12-15 cm

3-5 cm


7-10 cm


12-15 cm


No

of

pla

nts

/ m

2N

o o

f p

lan

ts /

m2

No

of

pla

nts

/ m

2N

o o

f p

lan

ts /

m2

0

10

20

30

40

50

60

70

BC

F

D

AB

F

DE

A

F

E

bc

d

ab abc

d

abc

d

abc

Pre application (15-May)

0

10

20

30

40

50

60

70

Figure 6. Effect of mowing heights and weed control treatments on seedling and mature plant densities of

dandelion after spring application (15 May 2004). S. minor rate�/60 g m�2; KillexTM herbicide rate�/1.7 kg

a.i./ha. Within a plant stage in each graph, bars labeled with similar letters are not significantly different at

P�/0.05 according to Tukey test.


Two weeks after S. minor treatment, the percentage of dandelion plants that

regenerated after complete foliar damage was significantly less in the spring (up to

15%) than in the fall (up to 32%) treatments. Although the mowing height did not

affect the percentage of root regrowth after the spring treatment of S. minor , there was

significantly less regrowth under the closest mowing height (22%) compared with the

two higher heights (33%) after the fall treatment (Figure 7).

Discussion

Previous studies on the virulence and efficacy of S. minor on dandelion have

demonstrated its biocontrol potential (Ciotola et al. 1991; Riddle et al. 1991; Briere

et al. 1992; Stewart-Wade et al. 2002b). Our results support those studies and

indicate the importance of correct mowing regimes on dandelion survival rates as

influenced by S. minor .

In the absence of weed control, periodic mowing at any of the studied levels was not

effective in controlling dandelion. However, compared to other heights, periodic

mowing at 5/5cm caused significant reduction in root biomass and field population

density of dandelion, but close mowing height caused a significant increase in

broadleaf groundcover percentage and diversity (M.H. Abu-Dieyeh & A.K. Watson,

unpublished). In another study, mowing every 2 weeks eliminated field bindweed but

did not prevent dandelion colonization (Timmons 1950). The high regenerative

capacity of dandelion roots (Stewart-Wade et al. 2002a) is the main cause of

recolonization.

The level of mowing height capable of exerting significant stress on plants is highly

variable and depends mainly on the plant species, the surrounding environment, the

time of the year, and the frequency of mowing (Zanoni et al. 1969; Meyer & Schmid

1999; Liu & Huang 2002; Narra et al. 2004). Although close mowing may be harmful

to weeds, it may also be harmful to turfgrass species resulting in increased weed

infestation by tipping the competitive balance in favor of the weeds (Busey 2003). The

major turfgrass species in our fields were Kentucky bluegrass and perennial ryegrass,

Spring

% o

f re

gro

wth

fro

m r

oo

ts

0

5

10

15

20

25

30

35

3-5 cm7-10 cm12-15 cm

Fall

aa

b

bcc c

Figure 7. Effect of mowing heights on regrowth of dandelion roots, 3-weeks post-treatment application.

Mowing heights were initiated 2 weeks prior to spring application and maintained throughout the

experiment. Bars with similar letters are not significantly different at the 5% level according to Tukey test.


and in well maintained turf, both species are recommended to be mowed at a medium

height, �/5�7 cm (Turgeon 1985; Fry & Huang 2004). Therefore, our grass mowing

regime at 5/5 cm may have impacted the growth and survival of dandelions resulting

in increased colonization by competitive species with more resilience and tolerance to

close mowing.

Under greenhouse conditions, the severe effect of the fungus on above and below

ground biomass of dandelion was apparent on the third day after application and was

complete within 2 weeks. Mowing height had no effect on the efficacy of S. minor in

the greenhouse experiment. This could be explained by the rapid destructive nature of

S. minor , a necrotrophic fungus causing wilting, collapse and death of the infected

plant parts (Abawi & Grogan 1979; Melzer et al. 1997). This rapid destruction is

achieved in high moisture conditions over a wide range of temperature (5�258C)

(Melzer & Boland 1994). Therefore, the variation among the microenvironments

within these mowing regimes likely did not affect S. minor performance.

In our field experiments, two dates were chosen, 15 May and 15 September, to

synchronize two factors; (1) suitable climatic condition for S. minor to cause disease,

and (2) high abundance of dandelion during periods of recruitment. Average daily

mean temperature, relative humidity (RH), and dew point for the 2-week post-

application period were 158C, 78% RH and 10.68C, respectively. Under these

conditions the fungus needs only 2�3 days to germinate, invade and colonize

dandelion plants. The maximum effect of the fungus occurs within 10�14 days and

then the fungus dies and disintegrates on the soil surface. Thus, the maximum effect

of S. minor on dandelion density was obtained 2 weeks after application. Foliar

turfgrass pathogens can cause more disease in microenvironments with higher

canopies (Fagerness & Yelverton 2001; Martin et al. 2001), but disease incidence of

lettuce, a major host of S. minor , was not affected by different microclimates of the

crop canopy (Melzer & Boland 1994).

By 2 weeks after application in the field, the fungus destroyed most of the dandelion

population in all treated plots without being influenced by changes in the

microenvironment under the different mowing heights. In the 3�5-cm mowing height

plots, bare soil was exposed within the thin grass canopy allowing greater sunlight

interception at the soil surface. Consequently, high dandelion seedling recruitment

occurred after the fungus lost viability and died. Full light is a major requirement for

germination of dandelion seeds (Letchamo & Gosselin 1996) and buried seeds are

unable to germinate (Noronha et al. 1997).

After the effect of S. minor declined, dandelion seeds started to germinate and

seedling emergence continued until the end of June, and then germination declined.

At mid-June, significantly greater seedling emergence occurred in plots with the 3�5-

cm mowing height compared with the 7�10- and 12�15-cm mowing heights.

Therefore, at 6 weeks post-application, population density of dandelion increased

significantly in the 3�5-cm mowing height plots compared with other mowing heights

and became similar to untreated plots. In plots treated with S. minor , dandelion

re-established from seed and survived better under the 3�5-cm mowing height than

under the 7�10- and 12�15-cm mowing heights. Consequently, dandelion biocontrol

with the fungus at the two higher mowing heights was as effective as the chemical

herbicide.

The initial effect of the herbicide on dandelion density was much slower than that of

the fungus, but by 6-weeks post-application the density was reduced by 90%,


significantly lower than S. minor effect. Mowing height did not interact with the

herbicide effect. Seedling recruitment after the herbicide treatment was very low, due

to persistence of the herbicide in the soil with a half-life of 2�269 days reported (Cox

1999). Treating dandelion at the reproductive stage with KillexTM reduced the

germination potential down to 4.8 and 18.4% after spring and fall treatments,

respectively (Abu-Dieyeh et al. 2005). Moreover the herbicide treatment significantly

reduced dandelion seed bank (10 cm depth) compared with the untreated, control

plots (M H. Abu-Dieyeh and A. K. Watson, unpublished).

Root regeneration of dandelion is one of its competitive features and makes control

a difficult task (Stewart-Wade et al. 2002a) since a small section of root can propagate

a new plant when covered by 5�10 cm of soil (Falkowski et al. 1989). Our greenhouse

results indicated that S. minor caused a highly significant reduction of root biomass,

but 20�50% of the treated dandelions had resprouted from the roots at 6 weeks post

application. Resprouted plants were very weak lacking vigour and characterized by

tiny roots and tiny leaf shoots. Thus, S. minor is not only attacking aboveground

biomass of dandelion but also affecting the roots, leaving them less likely to survive

grass competition and more prone to other biotic and abiotic stresses, especially

winter frost. Under greenhouse conditions, the closest mowing height caused

significant reduction of sprout percentage compared with the two higher mowing

heights (15 and 20 cm). There was no difference in the number of resprouted

dandelions obtained at 8 weeks compared with 6 weeks post-application. No signs of

disease development by S. minor were observed on these new shoots, but the weakness

of the root caused by the direct effect of the fungus on the mother plants may diminish

its survival under the additional stress of grass competition. The extensive defoliation

stress caused by repeated mowing at the 5-cm level reduced the root biomass

significantly in untreated plants and this may explain the reduction of regrowth under

this mowing height. Similarly in other perennial plants, repeated defoliation reduced

regrowth ability of Ranunculus acris L., after infection by the fungus S. sclerotiorum

(Green et al. 1998) and repeated cutting integrated with a rust fungus, Puccinia

punctiformis exerted synergistic control effects on growth rate and reproductive success

of Cirsium arvense (Kluth et al. 2003).

Mowing heights were initiated 2 weeks prior to the spring application, hence stress

on the roots would not be strong, whereas the accumulative effect of defoliation stress

by mowing at the 5-cm level over the growing season prior to the fall treatment

resulted in significantly less root regrowth compared with other heights. Dandelion,

allocates more resources for flowering and vegetative growth in the spring (Cyr et al.

1990) while nitrogen resources are restored in the roots at the end of summer

(Rutherford & Deacon 1974) which may explain the increase in the percentage of

regrowth in the fall. The maximum regrowth percentage reported in the field is lower

than greenhouse experiment which indicates that field environments exert more

ecological stress on dandelion leading to improved performance of S. minor .

In conclusion, in a low-maintained cool-season turf environment, integrating

S. minor with appropriate mowing could be as effective as a herbicide. Dandelion

suppression was the least under close mowing due to the new opened environment

which induced more germination from soil seed bank. Extensive periodic defoliation

by mowing and application of S. minor on flowering dandelion (spring application)

might be the cause of decreasing dandelion root survival through exhaustion of the

root carbohydrate and nitrogen reserves. Understanding these physiological changes


in dandelion roots under different mowing heights will support successful deployment

of S. minor for dandelion control. Additionally, monthly monitoring of the weed

species dynamics and turfgrass quality provides good portrayal of the biocontrol

system of interacting organisms and the turf environment.

Acknowledgements

The authors thank Philippe Seguin for his valuable help in statistical analysis and also

Miron Teshler for his help in field and technical work. The financial support from

Hashemite University, Zerka, Jordan and from the Natural Sciences and Engineering

Research Council of Canada (NSERC) Idea to Innovation (I2I) grant are gratefully

acknowledged.

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Increasing the Efficacy and Extending the Effective Application Period of aGranular Turf Bioherbicide by Covering with Jute Fabric

Mohammed H. Abu-Dieyeh and Alan K. Watson*

Progress in bioherbicide development has been hindered by the strict moisture and temperature requirements of the livingactive ingredient. Application of a jute fabric to areas treated with a Sclerotinia minor granular bioherbicide improvedbroadleaf weed control and broadened the effective application period to include the warm summer season. When turfgrassplots treated with the bioherbicide were covered with burlap fabric for 3 d, broadleaf weed (dandelion, white clover,broadleaf plantain, buckhorn plantain, ground ivy, and prostrate knotweed) control was greatly enhanced. The cover wasmade of natural jute fibers that retained water but had sufficient transparency to allow 33% light penetration for continuedgrowth of the grass. Virulence of the bioherbicide was maintained under elevated temperatures that would otherwise reduceefficacy. The bioherbicide was ineffective in the summer unless covered, but dandelion density, broadleaf weed groundcover, and dandelion survival were all reduced by the bioherbicide when plots were covered, even if applications were madein July. The efficacy of the bioherbicide was also enhanced under favorable conditions, and covering permitted reducedapplication rates without loss of efficacy. When applied at a rate of 20 g/m2 and covered, S. minor granules exertedsignificantly greater biocontrol of dandelion than 40 g/m2 without covering. Covering for up to 5 d did not cause anyadverse effects on the turfgrass. This approach may overcome one obstacle to the commercialization of the Sclerotinia minorbioherbicide, permitting its deployment under challenging environmental conditions.Nomenclature: Broadleaf plantain, Plantago major L. PLAMA; buckhorn plantain, Plantago lanceolata L. PLALA;dandelion, Taraxacum officinale G. H. Weber ex Wiggers TAROF; ground ivy, Glechoma hederacea L. GLEHE; prostrateknotweed, Polygonum aviculare L. POLAV; white clover, Trifolium repens L. TRFRE.Key words: Application technology, biological control, jute, SARRITOR, Sclerotinia minor, turfgrass.

Most turfgrass environments have weed problems andrequire a degree of management to be functional andaesthetically pleasing (Monaco et al. 2002). One hundredforty-five broadleaf species are classified as weeds in turfgrassenvironments (McCarty et al. 2001). A combination of two orthree herbicides is normally recommended to control a widespectrum of broadleaf weeds in turf (Emmons 1995).Pesticide use restriction and prohibition in urban environ-ments in many municipalities in Canada have intensified thesearch for biological control options. However, the bioherbi-cide approach has had limited commercial or practical successmainly because of problems in persistence and performanceunder suboptimal environmental conditions (Hallett 2005;Kennedy and Kremer 1996). Many potential bioherbicideshave shown inconsistent results when evaluated under fieldconditions. The absence of adequate moisture regimenscontinues to limit the development of many prospectivebioherbicides (Chittick and Auld 2001). Considerableresearch has been directed toward increasing moistureretention and improving bioherbicide formulation application(Daigle et al. 1990), including sodium alginate granules(Boyette and Walker 1985; Walker and Connick 1983),starch-based granules (Quimby et al. 1999), a modified pastaprocess (Connick et al. 1991), invert emulsions (Leathers et al.1993; Quimby et al. 1988), and hydrophilic polymers(Chittick and Auld 2001; Shabana et al. 1997). Additionally,

nutrients that promote rapid infection can reduce theimportance of a free-moisture or dew period constraint(Schisler et al. 1991).Dandelion is one of the most abundant weed species and is

considered a major weed in home lawns and grass amenities ofNorth America, Europe, and other countries (Abu-Dieyehand Watson 2007a,b; Larsen et al. 2004; Monaco et al. 2002).The fungus Sclerotinia minor Jagger (IMI 344141) has beenregistered as a biological herbicide (Sarritor)1 for dandelion inCanadian turfgrass (PMRA 2007). Sclerotinia minor is adestructive necrotrophic fungus causing collapse and death ofinfected plant parts (Abawi and Grogan 1979; Melzer et al.1997). This rapid destruction is achieved in high humidityconditions over a wide range of temperatures (5 to 25 C)(Melzer and Boland 1994). When applied to turfgrass, theeruptive mycelium of the S. minor bioherbicide does notpersist in the absence of a susceptible host and quickly decayswithin 10 d (Watson 2007). Sclerotinia minor is asporogenic,and the bioherbicide active ingredient does not spread beyondtreated areas.Turf field trials have confirmed the efficacy of S. minor in

controlling dandelion and reducing broadleaf weed groundcover (Abu-Dieyeh and Watson 2005, 2007a–c). The broadhost range of the pathogen has been well documented(Hollowell et al. 2003; Melzer et al. 1997) and 32 broadleafweeds, representing many of the common broadleaf weeds incool-season turfgrass environments, were susceptible to spotapplications (0.2 to 0.4 g/plant) of the bioherbicide (Abu-Dieyeh 2006). However, there was a high degree of variabilityin the mean foliar damage and the survival rate of broadleafweeds 3 wk after treatment. This variability was mainlyattributed to the different growth habits of weeds and the

DOI: 10.1614/WT-09-001.1* Research Assistant and Professor, Department of Plant Science, McGill

University, 21,111 Lakeshore Road, Ste-Anne-de-Bellevue, Quebec, Canada,H9X 3V9. Current address of first author: Department of Biology andBiotechnology, The Hashemite University, P.O. Box 150459, Zarqa, Jordan.Corresponding author’s E-mail: [email protected]

Weed Technology 2009 23:524–530

524 N Weed Technology 23, October–December 2009

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degree of contact between the bioherbicide granule and thehost plant tissues (Abu-Dieyeh 2006).Greenhouse and long-term field studies have shown the S.

minor bioherbicide to significantly reduce the population aswell as the aboveground and belowground biomass ofdandelion in turf to levels similar to those with the commonlyused turfgrass herbicides (Abu-Dieyeh and Watson 2005,2006, 2007a). When S. minor was combined with turfgrassoverseeding, dandelion populations were reduced by 70 to80% in the first year, increasing to 95% reduction in thesecond year (Abu-Dieyeh and Watson 2007a). The S. minorbioherbicide provided effective control of dandelion and otherbroadleaf weeds during the spring and early autumn whenmean daily temperatures ranged from 9 to 20 C, mean dailyrelative humidity (RH) was above 58%, and the dew pointranged from 3.6 to 17.4 C (Abu-Dieyeh and Watson 2007c).When applied outside these parameters, control efficacy wasgreatly reduced, thus limiting the effective application periodfor this bioherbicide.As conditions of high temperature and low moisture

prevailed during the summer, the pathological action of the S.minor formulation (Stewart-Wade et al. 2002), like all knownbioherbicides (Fravel 2005), was diminished or ceased. Forbioherbicides, environmental factors, such as temperature, freemoisture, and protection from ultraviolet irradiation, areoften critical in causing plant infection and disease (Hallett2005). The duration of leaf wetness and the ambienttemperature have a combined effect on infection (Zhangand Watson 1997). Moisture is an important requirement formany pathogens for inciting disease and is known to enhancethe pathogenicity of S. minor (Melzer and Boland 1994).In this work, we present and evaluate a novel method of

application of the S. minor bioherbicide in turfgrass that canovercome moisture and temperature constraints, improveweed control efficacy, and reduce the bioherbicide applicationrate.

Materials and Methods

Site Description. Field studies were conducted on theMacdonald Campus of McGill University, in Ste. Anne-de-Bellevue, QC, Canada (45u259N, 73u559W, 39.00 melevation). The soil was a St. Bernard sandy loam (9% coarsesand, 82% fine sand, 5% silt, 4% clay), with a pH of 6.6 and6.3% organic matter. The turfgrass area had received minimalmaintenance throughout its history, except for repeatedmowing during the growing season (May to October). Thegrass sward was approximately 90% Kentucky bluegrass (Poapratensis L.) and 10% creeping red fescue (Festuca rubra L.var. rubra s.l.). The lawn flora was highly diversified with 18broadleaf weed species observed throughout the study period,and the dominant weed species was dandelion. In spring, thelevel of infestation ranged from medium (40 to 60 dandelionplants/m2 and 30 to 60% grass ground cover) to severe (80 to120 dandelion plants/m2 and 10 to 20% grass cover). Whiteclover and broadleaf plantain were less abundant thandandelion but had high temporal and spatial frequency. Lesscommon to occasional broadleaf species included birdsfoottrefoil (Lotus corniculatus L.) common ragweed (Ambrosia

artemisiifolia L.), black medic (Medicago lupulina L.),Carolina false dandelion [Pyrrhopappus carolinianus (Walt.)DC], mouseear chickweed (Cerastium fontanum Baumg.),common vetch (Vicia sativa L.), and buckhorn plantain.Prostrate knotweed and ground ivy occurred as single weedspecies patches in compacted soil locations.

General Field Experimental Conditions. All field experi-ments were established in completely randomized designswith three or four replications per treatment, and eachexperiment was repeated once. Treatment plots were 0.1 m2

(20 cm by 50 cm) to 0.2 m2 (40 cm by 50 cm) withapproximately 80 cm alleys between plots. During the first3 d after application, plots were sprinkler-irrigated 2 h dailyto saturate the jute fabric. The wetted fabric floated on thelawn surface without compressing the turf. The fabricremained moist for a period of 4 to 6 h during the day.Assessments were conducted on the day before application,and weekly for 3 to 4 wk post application. The data werecalculated as percentage of pre-treatment values.

Bioherbicide Formulation. The fungus, S. minor (IMI344141), was isolated from diseased lettuce plants (Lactucasativa L.) from southwestern Quebec, and the stock culturewas maintained as sclerotia at 4 C. The mycelia of thegerminated sclerotia were used to inoculate autoclaved crackedbarley grain (1.4 to 2.0 mm in diameter) as described in Abu-Dieyeh and Watson (2006). The S. minor granularformulation was freshly prepared 2 wk before treatmentapplications with viability and virulence of the fungalinoculum confirmed before use (Abu-Dieyeh and Watson2006).

Jute Fabric Covering. Jute burlap is fabricated from twocultivated Tiliaceae species Corchorus capsularis L. andCorchorus olitorius L. (Saha and Sen 1992). A commercialhorticultural textile2 made from natural jute fiber was used tocover bioherbicide-treated plots in turf for 3 d to improvemoisture conditions. The fibers were approximately 1.0 mmthick and were cross-weaved in a 3 mm by 3 mm and 2 mmby 3 mm grid. One layer of the fabric decreased visible lightpenetration at the turfgrass surface by 66%, whereas twolayers of fabric reduced light penetration by 89%. For the July2007 experiments and during the 3 d of covering, the range oftemperatures recorded under the jute fabric was 16.4 to 35.8C, and the average temperature was 25.4 C, compared with arange of 15.7 to 32.7 C and an average temperature of 25.3 Cfor uncovered plots. The range of RH under the cover was 70to 100%, and the average was 94.5%, compared with a rangeof 51.3 to 100% and an average of 77.3% for uncoveredplots.

Effect of Bioherbicide Rate and Jute Fabric Covering onDandelion Density, Regrowth, and Broadleaf Cover.Spring Application. At application on May 20, 2005, thetemperature was 14 C, and the RH was 71.8%. Theexperiment included seven treatments: untreated and notcovered; untreated and covered by jute fabric for 3 consecutived; treated with 40 g/m2 of S. minor–inoculated barley grainand not covered with the jute fabric; treated with 10, 20, 30,or 40 g/m2 of S. minor–inoculated barley grain and covered

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with jute fabric for 3 consecutive d. The measured parameterswere number of dandelions per plot and percentage ofbroadleaf weed ground cover.

To monitor the regrowth of dandelions, 10 dandelionplants in each of the S. minor–treated plots were selected atrandom and marked using colored pins before treatment.Three wk after application, the number of marked dandelionplants that resprouted was recorded in each of the 40 g/m2

treated plots, and then a second application of S. minor–inoculated barley grain (40 g/m2) was applied with coveringconditions similar to the first application. The regrowth fromthe marked dandelion plants was counted after a further 2 wk.The entire experiment was repeated on May 20, 2006, whenthe temperature and RH at application were 22.6 C and52.3%, respectively.

Summer Application. The above experiment, established in thespring, was repeated on July 20, 2005, when the temperatureand RH were 29.9 C and 52.8%, respectively. The summerapplication was repeated in 2006 on July 20. The temperatureand RH were 29.5 C and 78.7% at application, the ambientmaximum temperatures remained at 30 to 32 C for 3 d ormore, and the RH dropped to 30% during the week ofapplication.

Effect of the Bioherbicide and Jute Fabric Covering onJuvenile Prostrate Knotweed. The experiment was conductedon May 10, 2006, in a turfgrass area heavily infested with anewly emerged, nonmowed prostrate knotweed population.Plants were less than 5 cm in height with 1.5 branches/plantand a main stem diameter of 0.75 mm. Three treatments(untreated, treated and covered, treated and not covered) werecompared. The temperature was 17.8 C and the RH 40%whenthe treated plots (0.1 m2) received 40 g/m2 of the S. minor–inoculated barley grains. Percentage of prostrate knotweedground cover was visually estimated for all plots 1 d before S.minor application, on day 3, and weekly for 3 wk afterapplication. The experiment was repeated the following year onMay 20, 2007, at a nearby location at the same site when thetemperature was 23.6 C and the RH was 55%.

Effect of the Bioherbicide and Jute Fabric Covering onEstablished Prostrate Knotweed, Broadleaf Plantain,Buckhorn Plantain, Ground Ivy, and White Clover. Totest the effect of combining the cover with lower applicationrates of S. minor on economically important turfgrass weeds,separate field experiments for each of these five weed specieswere established on June 17, 2006. At treatment applicationthe temperature was 25.2 C and the RH was 77.2%. Theexperiment was repeated on the same day nearby in the samelawn area. Experimental sites were selected based on thepresence of a target weed. The plot size was 0.2 m2 (40 cm by50 cm). Each experiment included eight treatments (untreat-ed and not covered; untreated and covered; treated at 20 g/m2

and not covered; treated at 30 g/m2 and not covered; treatedat 40 g/m2 and not covered; treated at 20 g/m2 and covered;treated at 30 g/m2 and covered; and treated at 40 gm2 andcovered). The percentage of prostrate knotweed, white clover,and ground ivy ground cover were visually estimated for allplots on the same day before S. minor application and thenweekly for 4 wk. The number of individual plants of

broadleaf plantain and buckhorn plantain was counted onthe day of application and then weekly for 4 wk.

Duration of Coverage and Thickness of the Jute Fabric.Field experiments were conducted in early summer (mid-June) and early autumn (late August) to determine thepossibility of reducing the number of days that the jutecovering remained on the plots.

Effect of Number of Days of Covering. A one-factor experimentwas established on July 17, 2007, in a lawn area infested withdandelion, broadleaf plantain, and white clover. The fourtreatments were no covering or 1, 2, or 3 d of covering. Thetemperature was 27.2 C and the RH was 53% when the S.minor–inoculated barley grain was broadcast-applied at a rateof 40 g/m2. The total percentage of broadleaf weed groundcover was estimated for all plots on the same day before S.minor application and then weekly for 3 wk. The experimentwas repeated on August 20, 2007, when the temperature atapplication was 19.8 C and the RH was 55.4%.

Effect of Number of Layers of the Jute Cover. To evaluate theeffect of number of folds or layers of the jute cloth covering onthe ability of S. minor to exert weed control, an experimentwas established on July 17, 2007. The temperature was 27.2 Cand the RH was 53% at application. Four treatments(untreated and not covered; treated and not covered; treatedand covered with a single layer of jute fabric; and treated andcovered with two layers of jute fabric) were compared. Plotsize, bioherbicide rate, and parameters measured were allsimilar to the above-mentioned experiment on the duration ofcovering. The experiment was repeated on August 20, 2007,when the temperature was 19.8 C and the RH was 55.4% atapplication.

Effect of Jute Fabric Covering on Turfgrass Biomass.Twenty-four potting trays (25 cm by 20 cm by 6 cm) werefilled with a mixture of pasteurized soil (sandy clay loam), sand,and commercial potty mix3 (2 : 1 : 1 v/v) and sown with acommercial grass seed mixture4 (30% Kentucky bluegrass,40% creeping red fescue [Festuca rubra L. var. rubra s.l.], and30% turf-type perennial ryegrass [Lolium perenne L.]) at 3 g/tray. The trays were placed in a greenhouse at 24 6 2 C with15 h of light/day at a photon flux density minimum of350 6 50 mmol/m2/s. After 3 wk of growth, 18 of the 24 potswere chosen to be used for the experiment based on similaritiesof grass vigor. The experiment was a completely randomizeddesign with six treatment levels and three replications and wasconducted twice, once on June 20, 2007, and repeated onAugust 15, 2007. Sclerotinia minor–inoculated barley grainswere applied at a rate of 3 g 60 g/m2 (the highestrecommended rate) on the surface of the premoistened soil.The trays were uncovered or covered for 3 or 5 d with two foldsof jute fabric. The trays were checked daily and misted withwater whenever needed. One week after the treatmentapplication, the aboveground grass biomass of each tray wasclipped with hedge shears5, whereas the belowground biomasswas left without irrigation for an additional 3 d, then the soilwith roots of each tray was compressed and rolled on a 2-mmmesh screen until all dry soil was removed. The roots were thencarefully washed with water and separated from the soil


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residues. The aboveground and belowground biomass of eachtray was separately placed in paper bags, oven dried at 80 6 1C for 72 h, and then weighed.

Statistical Analyses. Statistical analyses were performed usingthe SAS statistical package.6 To overcome the differences inthe dandelion density between plots of the same experiment,posttreatment data were adjusted as a percentage ofpretreatment data collected on the day of application. Becauseall experiments were spatially or temporally repeated, the datafrom both repeats were pooled, and then subjected to theLevene test of SAS to evaluate homogeneity of error variances.Except for experiments conducted to test the effect of numberof days of covering and number of layers of the jute cover, allother experimental data from both trials were combinedbecause error variances were homogeneous. Normality foreach parameter was tested on model residuals using theShapiro-Wilk test. The SAS GLM procedure of repeatedmeasures was used to determine the effect on weed densityand broadleaf weed ground cover through time. The maintreatment effect of other data (not time related) wasdetermined using one-way ANOVA for a completelyrandomized design. Tukey’s test (SAS) at P 5 0.05 was usedto separate the means for all analyses with significant effects,and the paired t test was used to compare dandelion shootregrowth between the first and the second applications of thesame treatment.

Results and Discussion

Jute Covering Promotes the Efficacy of the Bioherbicide inthe Spring. Previous research showed that 60 g/m2 of the S.minor–inoculated barley grain applied in the spring or the fallwas required for acceptable dandelion control in low-maintained turfgrass (Abu-Dieyeh and Watson 2006,2007c) or 40 g/m2 if the bioherbicide was combined withturfgrass overseeding at 0 or 10 d after application (Abu-Dieyeh and Watson 2007b). In the spring, when plots werecovered with jute fabric for 3 d, a bioherbicide rate as low as20 g/m2 was enough to exert better control of dandelion than40 g/m2 without the cover (Figure 1A). Effective control onother broadleaf weeds, including white clover, broadleafplantain, and several other common weeds, was also achievedwith the low rate (Figure 1B). When covered with jute fabric,the efficacy of 20, 30, and 40 g m22 rates of the S. minorbioherbicide were the same (Figure 1). Although there wereno significant differences in dandelion regrowth from rootsbetween covered and uncovered plots 3 wk after the firstapplication, greater reductions in regrowth were noted afterthe second application in covered, compared with uncovered,plots (Figure 1C). These results indicate a better biocontroleffect when the treated plots are covered with jute fabric.Massive quantities of white mycelia were observed on thesecond day after application when plots were covered.

Jute Covering Broadens Bioherbicide Application Win-dow. In this study, the summer applications (20 July 2005and 2006) of 40 g/m2 of S. minor without cover were noteffective in controlling dandelions and other broadleaf weedseven though all study plots received 2 h of sprinkler irrigation

for 3 d after treatment (Figure 2). During the first 2 wk afterapplication, the recorded temperatures were 11 to 32.8 C, theRHs 27 to 99%, and the dew points 9.5 to 23. The averagemaximum daily temperature was 29.5 C, and the averageminimum daily RH was 37%.

Summer climate conditions are generally not suitable forthe S. minor bioherbicide to cause disease, but when thetreated plots were covered with jute fabric, good control wasobtained with rates as low as 20 g/m2, and better control wasachieved with 30 and 40 g/m2 (Figures 2A and 2B). WhenRH was maintained above 90% under the cover during thefirst 3 d after application, the fungus continued rapid growtheven though the temperature under the cover increased to 35C. This indicates the importance of RH, not temperature, toexplain poor control in uncovered plots during the summer.The RH under the cover was 94.5% (70 to 100%) comparedwith 77.3% (51.3 to 100%) for uncovered plots. The jutefabric cover retains moisture and preserves free moistureaccumulated on the soil and the leaves from sprinkler wateringor natural dew. Moreover, the jute cover presumably reducesUV exposure and helps to protect the fungus from UV light.Although inadequate moisture is the most commonly citedreason for failure of bioherbicides when applied under fieldconditions, UV light could be the cause behind the failure ofcertain bioherbicides (Hallett 2005). In this study, dandelionsurvival, expressed as shoot regrowth from perennial root

Figure 1. Effect of Sclerotinia minor applied in May and then covered by jutefabric or uncovered on (A) dandelion density, (B) broadleaf weed ground cover,and (C) root regrowth of dandelion. The data in graph A and B are based on oneS. minor application performed on May 20, 2005. In graph C, the secondapplication was done 3 wk after the first application. Regrowth data wereobtained 2 to 3 wk after each application. Within each time assessment orapplication, means with a common letter are not significantly different atP 5 0.05 according to Tukey’s test. * refers to significance at P 5 0.05 betweenthe two applications of the same treatment.


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stocks, was 28% after the first application and only 15% afterthe second application of 40 g/m2 (Figure 2C).

Jute Cover Enhances Control of Weeds with DifferentGrowth Habits. In the early spring study, a significantbiocontrol effect of S. minor at 40 g/m2 rate occurred onnewly emerged prostrate knotweed when covered by jute, butcontrol was delayed in the uncovered and treated plots andonly reached 60% at 14 d after application (Figure 3A).Greater control during the summer with established prostrateknotweed was also observed with covered, compared withuncovered, plots (Figure 3B). Likewise, the jute coverpromoted consistent control of white clover, broadleafplantain and ground ivy, where control was better under thejute cover even when half the bioherbicide rate was applied(Figure 4). The least controlled species was buckhornplantain. The upright habit and lanceolate leaves of buckhornplantain may contribute to reduced contact with thebioherbicide product and thereby contribute to the decreasedlevels of control observed. Without the jute cover, controlremained poor even 7 d after application. However, 60 to70% control of buckhorn plantain was obtained with 40 g/m2

of S. minor granules when covered with jute (Figure 4B).Jute cover consistently increased the efficacy of the S. minor

bioherbicide on all the studied weed species. The jute coverpositively modified the microenvironment to provide optimal

conditions for S. minor mycelia growth, which dispersed themycelia throughout the entire plot, enabling greater contactwith host plant tissues and plant death.

Duration of Coverage and Number of Folds of theJute Fabric. One day of covering was not effective in mid-July or even in late August (Figure 5). Two days of jutecovering provided up to 90 to 100% suppression of broadleafweeds during late August application, but 3 d of jute coveringwere needed to obtain the same level of control during mid-July (Figure 5). Folding the jute cloth to have two layers,instead of one, significantly increased the efficacy of S. minorin controlling broadleaf weeds, especially in the summer(Figure 6). We presume that two folds of jute fabric retainedmore water for a longer period and more effectively promotedS. minor growth, particularly during summer conditions.Under favorable conditions, the fungus S. minor needs 3 to5 d to reach its optimal growth and to cause extensive damageon broadleaf weeds (Abu-Dieyeh, 2006). Using two folds ofjute may reduce the amount of time the plots need to becovered during spring and fall conditions, but this would notbe possible during summer conditions because removing thecover before 3 d would result in killing the fungus, ceasing itsactivity.

Turfgrass Was Not Affected by Covering with Jute. Duringall the above field experiments, no signs of weakness, damage,or disease symptoms were observed on the turfgrass in any ofthe treated and jute-covered plots. Moreover, greenhouseexperiments confirmed that the belowground, aboveground,and total biomass of turfgrass was not affected after 3 or 5 d ofcovering with jute (data not presented). Field and greenhousestudies have confirmed that S. minor IMI 344141 is notpathogenic to turfgrass species (Abu-Dieyeh and Watson2007b). Members of the grass family Poaceae are notsusceptible to S. minor (Melzer et al. 1997).

Feasibility and Practicality of Using Jute Cover witha Bioherbicide. Fibrous barriers are commonly used innursery, horticultural, and landscaping to exclude all plantgrowth (Derr 1989; Feldman et al. 2000; Forcella et al. 2003;

Figure 2. Effect of Sclerotinia minor applied in July and then covered by jutefabric or uncovered on (A) dandelion density, (B) broadleaf weed ground cover,and (C) root regrowth of dandelion. The data in graph A and B are based on oneS. minor application performed on May 20, 2005. In graph C, the secondapplication was done 3 wk after the first application. Regrowth data wereobtained 2 to 3 wk after each application. Within each time assessment orapplication, means with a common letter are not significantly different atP 5 0.05 according to Tukey’s test. * refers to significance at P 5 0.05 betweenthe two applications of the same treatment.

Figure 3. Effect of Sclerotinia minor on prostrate knotweed when covered by jutefabric or uncovered. (A) Juvenile knotweed (beginning of May trial). (B)Established knotweed (mid-June trial). Within each time assessment, means witha common letter are not significantly different at P 5 0.05 according toTukey’s test.


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Martin et al. 1991). Covers and grass seed blankets are oftenused to enhance plant germination and turfgrass establishmentrates by retaining soil moisture and increasing soil tempera-ture (Miltner et al. 2004; Patton et al. 2008). These examplesof the use of covers provide economic advantage and benefitfor home owners and other users. Thus, procedures such asthis are not without precedent. Nonetheless, the applicationand removal of the fabric is an additional activity forcommercial applicators or home owners and may present asignificant barrier to adoption. The covering technique wastested over a range of environmental conditions, and it always

exerted improved biocontrol results compared with theuncovered–treated plots. The cover lowered use rates andbroadened the application window to include warmer anddrier conditions. The above advantages may be applicable toother bioherbicide projects in other ecosystems and addressmany of the obstacles that have hindered the development ofmany bioherbicides.

The present cover is made of natural jute fibers. The fibermat is readily available, easily applied, storable, reusable,relatively inexpensive (approximately $2 per 100 m2) andenvironmentally safe. It seems likely that the jute fabric couldbe replaced by sheets made of any number of other naturalplant fibers or combinations of plant fibers. Synthetic fibers,such as polyester, polyethylene, and polypropylene, may alsobe effective, but further testing for effectiveness and absence ofturf damage would be required.

Sources of Materials1 Sarritor Biological Herbicide, Sarritor Inc., Notre-Dame-de-

l’Ile-Perrot, QC, Canada, J7V 8P1.2 Jute fabric, TerraTex, Lenrod Industries Ltd, Ville St-Laurent,

QC, Canada, H4N 1V8.3 Promix soil, Promix BXt, Premier Horticulture Ltee., 1

Premier Avenue, Riviere-du-Loup, QC, Canada, G5R 6C1.4 Commercial turfgrass seeds, C.I.L. Golfgreen, Spectrum Brands

Canada Ltd., 10 Craig St., Brantford, ON, Canada, N3R 7J1.5 Hedge shears, PlantSmart, WalMart, 17101 Ch. Sainte-Marie,

Kirkland, QC, Canada, H9J 1M3.6 SAS Statistical Package, 2002 Edition, SAS Institute, Cary,

NC, 2002.

Acknowledgments

The authors thank Inaam Shaheen and Miron Teshler fortheir help in laboratory and field work. The financial supportfrom the Natural Sciences and Engineering Research Councilof Canada (NSERC) Idea to Innovation (I2I) grant isgratefully acknowledged.

Figure 4. Effect of Sclerotinia minor on (A) broadleaf plantain, (B) buckhornplantain, (C) white clover, and (D) ground ivy densities in turf grassenvironments when covered by jute fabric or uncovered. Within each timeassessment, means with a common letter are not significantly different atP 5 0.05 according to Tukey’s test.

Figure 5. Effect of Sclerotinia minor on broadleaf weed ground cover whencovered with jute fabric for 1, 2, or 3 d or uncovered in a (A) mid-June trial and(B) late August trial. Within each time assessment, means with a common letterare not significantly different at P 5 0.05 according to Tukey’s test.

Figure 6. Effect of Sclerotinia minor on broadleaf ground cover when covered byone or two folds of jute fabric or uncovered in a (A) mid-June trial and (B) lateAugust trial. Within each time assessment, means with a common letter are notsignificantly different at P 5 0.05 according to Tukey’s test.


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Received January 4, 2009, and approved June 23, 2009.


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RESEARCH ARTICLE

Development of strain specific molecular markers for the Sclerotiniaminor bioherbicide strain IMI 344141

L. Pana$, G.J. Ashb, B. Ahna and A.K. Watsona*

aDepartment of Plant Science, McGill University, 21,111 Lakeshore Road, Ste-Anne-de-Bellevue, Quebec, Canada H9X 3V9; bE.H. Graham Centre for Agricultural Innovation, Charles

Sturt University, Locked Bag 588, Wagga Wagga, NSW 2678, Australia

(Received 13 April 2010; accepted 4 May 2010)

The plant pathogenic fungus, Sclerotinia minor IMI 344141, has been developedas a bioherbicide for broadleaf weed control in turfgrass and a means todifferentiate this biocontrol agent from like organisms is required. A strainspecific molecular marker was developed to detect and monitor the Sclerotiniaminor IMI 344141 bioherbicide strain. The method was based on polymerasechain reaction (PCR) amplification of two sequence-characterized amplifiedregions (SCAR) primer pairs for a first round PCR, and another two sets ofnested primers was used for a second round PCR if higher sensitivity was needed.Sclerotinia minor IMI 344141 was successfully traced from both pure cultures andenvironmental samples originating from bioherbicide-released field trials. DNAof the S. minor bioherbicide isolate IMI 344141 was detected in the soil 2 monthsafter application, but was not detected in the 3- and 9-month samples afterapplication. When applied as a bioherbicide, S. minor (IMI 344141) did notpersist into the following spring season in turf environments. This moleculardetection method provides a mechanism to distinguish this isolate from relatedorganisms and a tool to monitor behavior of the biocontrol agent S. minor IMI344141 in nature, particularly in soil.

Keywords: biocontrol; bioherbicide; detection; mycoherbicide; PCR; RAPD;SCAR; Sclerotinia minor; turfgrass

Introduction

Dandelion (Taraxacum officinale Weber ex Wiggers) represents one of the single

largest target pests for application of pesticides in North America. Chemical control

has been the accepted method of dandelion and broadleaf weed control in turfgrass,

with an estimated North American market of $300 million US$ per year. Growing

awareness of environmental and human health concerns led numerous municipal and

several provincial governments in Canada to ban or severely restrict use in urban

settings, leaving limited options for effective weed control in turfgrass. A biological

option is now available with the commercialization of the SarritorTM bioherbicide

based on Sclerotinia minor (PMRA 2007; Abu-Dieyeh and Watson 2007c).

*Corresponding author. Email: [email protected],$When originally published online the author name was incorrect. The online version has nowbeen amended.

Biocontrol Science and Technology,

Vol. 20, No. 9, 2010, 939�959

ISSN 0958-3157 print/ISSN 1360-0478 online

# 2010 Taylor & Francis

DOI: 10.1080/09583157.2010.491895

http://www.informaworld.com

Downloaded By: [Canadian Research Knowledge Network] At: 17:15 22 November 2010

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Sclerotinia minor Jaggar is a soil borne Discomycetes fungus characterized by

small (0.5�2.0 mm), irregular sclerotia that germinate by eruptive growth of

mycelium. Apothecia are rarely observed under field conditions and sclerotia are

the main means of S. minor survival and dispersal. Sclerotinia minor is a broad

spectrum, destructive, necrotrophic fungus that causes collapse and death of infected

plants, and plant parts (Abawi and Grogan 1979). The broad host range of S. minor

has been well documented with significant crop losses recorded in lettuce andpeanuts (Melzer, Smith and Boland 1997; Hollowell, Shaw, Cubeta and Wilcut 2003).

One isolate of S. minor, IMI 344141 collected in south-western Quebec from diseased

lettuce, has been registered as a biological herbicide (SarritorTM) for control of

dandelion and other broadleaf weeds in Canadian turf grass environments (PMRA

2007). The efficacy of S. minor IMI 344141 in controlling dandelion and reducing

broadleaf weed ground cover have been confirmed in numerous field trials (Abu-

Dieyeh and Watson 2006, 2007a,b,c).

Microbial pest control agents (MPCA) must be characterized to show relatedness

to like organisms and to distinguish the MPCA from related microorganisms. From

critical regulatory, commercial and litigation perspectives, methodologies that enable

the IMI 344141 strain ofS.minor to be distinguished from like-organisms are essential.

Traditionally, Sclerotinia species are identified by morphological and physiologi-

cal criteria such as gross cultural characteristics, sclerotial size, ascus and ascospore

dimensions, etc. These methods are still used and sclerotial diameter remains a goodcriterion for separation of S. minor from S. sclerotiorum and S. trifoliorum (Ekins,

Aitken and Coulter 2005). However, there is no difference in the structure of hyphae

among S. sclerotiorum, S. trifoliorum, and S.minor. The high morphological similarity

among Sclerotinia species may make it difficult distinguishing S. minor from others

(Willetts and Wong 1980). These traditional identification methods involve tedious

and time-consuming work, particularly to distinguish one isolate from others.

Phenotypic variation among 30 isolates ofS.minor, collected fromdifferent regions

and hosts, was studied in an attempt to characterize the IMI 344141 strain (Shaheen,

Abu-Dieyeh, Ash and Watson 2010). There was high diversity (0.6) among mycelial

compatibilitygroups (MCG) and isolates varied significantly in sclerotia shape (length/

width ratio) and number, but did not vary in colony morphology or growth. IMI

344141 could be phenotypically distinguish from the other S. minor isolates examined

by performing vegetative compatibility testing and enumeration of sclerotia produced

on standard potato dextrose agar (PDA) plates. But without molecular marker

genotyping, MCGs and phenotyping do not suffice to completely characterize and

distinguish the IMI 344141 strain from broader S. minor populations.DNA-based biotechnological methods have become useful in detection of disease

and tracking of organisms in nature. In particular, the PCR amplification method

has been useful by tracking specific marker DNA sequences that are designed based

on sequences of an organism or genes that need to be monitored in nature, for

instance a genetically modified organism (Jansson 1995). The technique, called

sequence characterized amplified regions (SCAR) technique, is now widely used in

all areas of biology including detection of a biocontrol agent after application in the

field (Castrillo, Vandenberg and Wraight 2003; Dauch, Watson and Jabaji-Hare

2003; Pujol, Badosa, Cabrefiga and Montesinos 2005; Nunes et al. 2008). A strain

specific marker was developed to detect the biocontrol agent, Colletotrichum

coccodes, a prospective bioherbicide for velvetleaf (Abutilon theophrasti), that

940 P. Li et al.


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successfully identified C. coccodes in natural samples (Dauch et al. 2003). Biocontrol

molecular markers can be developed using sequence polymorphisms in internal

transcribed spacer (ITS) sequence of nuclear ribosomal DNA (Freeman et al. 2002),

randomly amplified polymorphic DNA markers (Dauch, Watson, Seguin and Jabaji-Hare 2006), inter simple sequence repeat (ISSR) markers, RFLP, AFLP, etc.

However, the RAPD technique is popular because its procedure is straightforward

compared to other methods, and has minimal equipment requirement. The objective

of this study was to develop a method to detect and identify S. minor IMI 344141

from natural samples.


Biological material and growth conditions

Thirty isolates of Sclerotinia minor (Table 1) and 24 heterogeneous organisms (Table 2)

used in this study were collected from various hosts and from diverse localities. Stock

cultures of the fungal isolates were prepared on PDA (Difco Laboratories, Detroit,

MI, USA) and stored on PDA under sterile mineral oil at 48C.All fungal organisms were grown on PDA at 208C in the dark for 4�5 days. Five

agar plugs from the edge of actively growing colonies were transferred to potatodextrose broth (PDB, Difco Laboratories). Cultures were incubated on a rotary

shaker at 200 rpm at 20�248C for 5 days. The cultures were then filtered through

Whatman No. 1 filter paper (Fisher Scientific, Nepean, ON, Canada) and mycelia

were harvested. The mycelia were lyophilized and kept at �808C until DNA

extraction.

Bacteria were grown in 20 mL of nutrient broth media (Becton-Dickinson

Microbiology Systems, Cockeysville, MD, USA) at 100 rpm for 48 h. The bacterial

culture was centrifuged at 13,000 g for 2 min and the pellet was directly used forDNA extraction.

Plants were grown in pots containing pasteurized greenhouse soil and placed in a

greenhouse at 24/208C day/night. Plants were cut above ground level and kept at

�808C until they were used for DNA extraction. The S. minor IMI 344141 granular

formulation used for field application were produced according to Abu-Dieyeh and

Watson (2006).

Preparation of nucleic acids

Fungal genomic DNAwas extracted from 20 mg lyophilized mycelia according to Lee

and Taylor (1990). Plant DNA was extracted with DNeasy Plant mini kit (Qiagen,

Mississauga, ON, Canada) according to the manufacturer’s instruction. Bacterial

DNAwas extracted according to Wilson (2001). DNA isolated from fungi, bacteria,

and plants was quantified by Nanodrop ND-1000 (Nanodrop Technologies,

Wilmington, DE, USA) and adjusted to 5 ng/mL with 10 mM Tris�HCl (pH 7.6).DNA extraction from soil was performed with the UltraClean Soil DNA

Isolation kit (MoBio Laboratories, Solana Beach, CA, USA) following the

manufacturer’s instructions, except using a FastPrep apparatus (Thermo Savant,

Holbrook, NY, USA) at a speed 4 for 40 s instead of vortexing to dislodge DNA

from soil.

Biocontrol Science and Technology 941


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RAPD-PCR procedure

Three hundred Operon decamer primers and 100 UBC microsatellite primers were

used for initial screening with a subset of three to six S. minor isolates and several

heterogeneous organisms. One strain of S. minor from IMI 344141’s (isolate 13)

vegetative group (isolates 7, 10, 13, 15, 22, and 28) (Shaheen et al. 2010), in addition

to 13, was always included in the initial screening. The sets of Operon decamer

primers used in the initial screening were OPA, OPB, OPC, OPE, OPF, OPH, OPJ,

OPK, OPN, OPQ, OPS, OPAA, OPAB, OPAM, and OPBA (Operon Technologies,

Huntsville, AL, USA), 20 primers per set. The microsatellite primers (Inter simple

sequence repeat: ISSR) set #9, UBC801-UBC900, were used.

The RAPD PCRwas performed in 25 mL of the reaction mix consisting of 10 ng

DNA template, 20 mM Tris�HCl (pH 8.4), 20 mM KCL, 1.5 mM MgCl2, 200 mM of

each dNTP, 500 nM primers, and 1 U of Taq polymerase (Invitrogen, Carlsbad, CA,

USA). Amplification reactions were carried out in a GeneAmp PCR system 9700

Table 1. Sclerotinia minor isolates used in this study.

Isolate code/

number

Electrophoresis

designation Host Origin

R21 1 Peanut (Arachis hypogaea) Virginia, USA

R22 2 Lettuce (Lactuca sativa) New Jersey, USA

R23 3 Peanut (Arachis hypogaea) Virginia, USA

R24 4 Endive (Cichorium endivia) New Jersey, USA

R25 5 Cabagge (Brassica campestris) New Jersey, USA

LRC2104 6 Lettuce (Lactuca sativa) Manitoba, Canada

JW1 7 Unknown Hull, UK

ATCC44236 8 Lettuce (Lactuca sativa) New York USA

SMRF02 9 Peanut (Arachis hypogaea) Virginia, USA

BRIP28139 10 Chickpea (Cicer arientium) New South Wales, AU

Sm44 11 Lettuce (Lactuca sativa) California, USA

Sm66 12 Lettuce (Lactuca sativa) Ontario, Canada

IMI 344141 13 Lettuce (Lactuca sativa) Quebec, Canada

TH1C 14 Canola (Brassica napus) New South Wales, AU

VPRI1671 15 Onion (Allium sativum) Tasmania, AU

VPRI1285 16 Bathurst burr (Xanthium spinosum) Victoria, AU

BPIC1949 17 Lettuce (Lactuca sativa) Kifissia, Greece

LU286 18 Lettuce (Lactuca sativa) Canterbury, NZ

LK1 19 Centaurea pannonica Budapest, HU

MUCL38484 20 Unknown Belgium

MUCL11551 21 Lettuce (Lactuca sativa) Belgium

K4205 22 Mungbean (Vigna radiata) Queensland, AU

S94-001 23 Unknown Suwon, Korea





CBS112.17 28 Lettuce (Lactuca sativa) Netherlands

CBS207.25 29 Sasify (Tragopogon porrifolius) Netherlands

CBS339.39 30 Lettuce (Lactuca sativa) Italy

942 P. Li et al.


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(Applied Biosystems, Streetsville, ON, Canada) for a denaturation step (3 min at

948C), 45 cycles of amplification (1 min at 948C, 1 min at 378C, 2 min at 728C), andfollowed by a final extension step (10 min at 728C). A PCR negative control in which

H2O replaced DNA was included in each PCR run.

Only primers that generated interesting polymorphic patterns of amplified

products between IMI 344141 and other isolates were retained for a second round

screening with all homogeneous and heterogeneous organisms. Thirty-five decamer

primers and one ISSR primer were chosen for the second round screening. Among

these primers, DNA fragments amplified by OPBA14, OPQ13, OPJ07, OPAA15

Table 2. Heterogeneous species used in this study.

Isolate code/

number

Electrophoresis

designation Host Origin

S. sclerotiorum WM-A1 31 Unknown Washington,

USA

S. sclerotiorum Lu457 34 Giant buttercup Canterbury, NZ

S. sclerotiorum Lu460 35 Tomato Canterbury, NZ

S. sclerotiorum Lu461 36 Scotch thistle Canterbury, NZ

S. sclerotiorum Lu462 37 Kale Canterbury, NZ

S. sclerotiorum Lu481 38 Lactuca sativa Canterbury, NZ

S. sclerotiorum KAC 39 Peach California, USA

S. sclerotiorum SS26A 51 Lettuce- Sherrington

Quebec, Canada

S. trifoliorum L-1 52 Red clover Mississippi,

USA

S. trifoliorum L-2 53 Red clover Mississippi,

USA

S. trifoliorum 3-5 56 Alfalfa Mississippi,

USA

S. trifoliorum HV-5 57 Hairy vetch Mississippi,

USA

Monilinia laxa STE-U 5724 59 Nectarine-Flamekist Matieland,

South Africa

M. laxa VPRI 163 60 Unknown Knoxville, AU

M. fructicola VPRI 1907 61 Unknown Knoxville, AU

M. fructicola 33-B11 62 Unknown Knoxville, AU

M. fructicola 33-E9 63 Unknown California, USA

Sclerorium cepivorum S.c. 01-01 64 Unknown California, USA

Botrytis fabae sardina VPRI 1758 65 Unknown Washington,

USA

Xanthomonas

campestris f.sp. Xep PA2 B1 NA NA

phaseoli

Xanthomonas

campestris f.sp. Xep STD3 B2 NA NA

phaseoli

Annual bluegrass A NA Quebec, Canada

Perennial ryegrass P NA Quebec, Canada

Dandelion D NA Quebec, Canada



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RAPD primers and UBC846 were further analyzed for sequencing due to their

promising aspect for amplifying a unique amplicon in the S. minor bioherbicide

strain.

Sequencing and cluster analysis

The target amplicons were gel-purified by Qiagen Gel Extraction kit (Qiagen) and

cloned with TA cloning kit (Invitrogen). Plasmids were used to transform E. coli

competent cells (One Shot TOP10, Invitrogen) according to the manufacturer’s

protocol.

Plasmid DNA was extracted according to Sambrook, Fritsch and Maniatis

(1989) and sent to McGill University and Genome Quebec Innovation Centre

(Montreal, QC, Canada) for sequencing. The DNA fragment sequences wereanalyzed with clustal W multiple sequence alignment software (version 3.2) at the

Biology Workbench website (http://workbench.sdsc.edu/). Sequences of the ampli-

cons were compared to nucleotide sequences of other organisms using BLASTN to

check potential homologies of the sequences.

SCAR (sequence characterized amplified regions) primer design and PCR conditions

According to the sequences of amplicons produced by OPBA14, OPQ13, OPJ07, and

OPAA15 RAPD primers, sets of SCAR primers with 18�24 bp were designed using

the Primer3 software at the Biology Workbench website. SCAR primers were firsttested for their specificity against six isolates of S. minor. Only those that highlighted

few isolates were selected for second round screening against all the S. minor isolates,

then third round screening was against heterogeneous organisms.

One SCAR primer set of OPBA14 (OPBA14-52/31) and one primer set of OPJ07

(J7L6/R1) were selected to discriminate isolate 13 from all organisms (Table 3). In

Table 3. RAPD, SCAR, nested PCR and ITS PCR primers used in this study.

Primer Sequence (from 5? to 3?)Length

(bp)

Tm

(8C)Amplicon

size (bp)

RAPD OPJ-07 CCTCTCGACA 10 37 737

OPBA-14 TCGGGAGTGG 10 37 1423

SCAR OPJ7-L6 CCTCTCGACAACCGACTAAAA 21 60 669

OPJ7-R1 GGGTTGCATAAGGTAAGCGA 20 60

BA14-52 GCCTGGCTGGTCGTAGCG 18 60 1414

BA14-31 TCGGGAGTGGGAATGGGGAT 20 60

28-QR GTATACTATGGTCGTGGT 18 55 900 or 400

Nested JF2 GCATTGCTAGCGTTGTAGTTGC 22 60 280

JR2 AATGAAGCTGTGGAAGGGAGAG 22 60

BA53 TGGCTGGTCGTAGCGATG 18 60 1392

BA33 TGGGGATGGGGATGGGAA 18 60

ITS ITS-1F CTTGGTCATTTAGAGGAAGTAA 22 60 �530

ITS-4 TCCTCCGCTTATTGATATGC 20

944 P. Li et al.


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order to increase sensitivity of the PCR, sets of nested primers were designed from

OPBA14 and OPJ07 sequences. Nested PCR primers were also tested for their

specificity against all homogeneous and heterogeneous organisms.

The SCAR PCRwas performed in 25 mL of the reaction mix consisting of 10 ng

DNA template, 20 mM Tris�HCl (pH 8.4), 20 mM KCl, 1.5 mM MgCl2, 200 mM of

each dNTP, 500 nM primers, and 1 U of Taq polymerase (Bioshop Canada Inc.,

Burlington, ON, Canada). Amplification reactions were carried out in a GeneAmpPCR system 9700 (Applied Biosystems) for 30 cycles of amplification steps (948C for

30 s, 608C for 30 s, 728C for 1.5 min for OPBA14 and 1 min for OPJ07) after an

initial denature step (3 min at 948C), and was finalized by an extension step at 728Cfor 10 min. A PCR negative control in which H2O replaced genomic DNA was

included in each PCR run. For nested PCR, the condition was the same except that

20� diluted products from first round PCR were used and extension time for JF2/

R2 was 30s at 728C for 30 cycles. In order to insure the quality and integrity of the

extracted DNA and the presence of fungal DNA in the soil sample to reduce the risk

of obtaining a false-negative result, DNA samples extracted from soil were amplified

with ITS-1F/ITS4 primers that recognize the internal transcribed spacers (ITS1-

ITS2) of fungal nuclear ribosomal DNA (White, Bruns, Lee and Taylor 1990).

Electrophoresis and analysis of amplification products

The PCR products were analyzed by electrophoresis on 1.2% agarose gels in 1�TAE buffer. 100 bp or 1 kb Plus DNA ladder (Invitrogen) or Lambda DNA/Hind III

Plus ladder (Bioshop Canada Inc.) was used as a molecular weight marker. Gels were

stained with ethidium bromide and images were recorded by BIO-RAD Quantity

One 4.2.1.

Sclerotinia minor IMI 344141 detection on leaves of lettuce and dandelion

Romaine lettuce and dandelion plants were grown from seeds in a greenhouse for

2 months. Isolates 13 and 28 were grown on PDA from sclerotia at 208C in the dark

for 4�5 days. Mycelia plugs of each isolate (8 mm in diameter) from the edge of

actively growing colonies were transferred onto leaves of potted lettuce and

dandelion. Three plugs of isolate 13, isolate 28, or both isolates were placed on

two leaves each of lettuce and dandelion. After 24 h, leaves were cut, immediatelyfrozen in liquid nitrogen, and kept at�808C until they were used for DNA

extraction. This experiment was repeated as above but leaves were harvested after

4 days in the repeat experiment in order to obtain more fungal colonization of the

leaves.

Genomic DNA was extracted from 100 mg of lyophilized leaves using DNeasy

Plant mini kit. The PCR was performed in 50 mL of the reaction mix consisting of

1 ng of DNA template, 20 mM Tris�HCl (pH 8.4), 20 mM KCl, 1.5 mM MgCl2,

200 mM of each dNTP, 500 nM primers (OPBA-52/28-QR), and 1 U of Taq

polymerase (Bioshop Canada Inc.). The PCR program included: an initial denature

step (3 min at 948C), 30 cycles of amplification steps (948C for 30 s, 528C for 30 s, 728Cfor 1 min), and a final extension step at 728C for 10 min. The positive control was

mycelia DNA of isolate 13. The negative control was H2O replaced genomic DNA.



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Detection of Sclerotinia minor IMI 344141 sclerotia in soil

Soil was collected from a control field never exposed to S. minor. One flask of soil

was autoclaved twice on two consecutive days for 1 h at 1218C and 15 p.s.i. Two-

tenths gram of soil was measured for each of three tubes of autoclaved soil and three

tubes of field soil. Average sized sclerotia were selected at random and one sclerotium

was added into each tube with the soil. Tubes were placed in a growth chamber at

208C in dark. After 15 days, tubes were frozen in liquid nitrogen and stored

at�808C until extraction.

Genomic DNA was extracted using the UltraClean Soil DNA Isolation kit

(MoBio Laboratories, Solana Beach, CA, USA) as described above, except it was

preceded by grinding of the 0.2 g of soil containing a sclerotium. PCRwas performed

with SCAR primers BA14-52/31, OPJ7-L6/R1 and ITS-1F/ITS4 as described above

in SCAR PCR conditions, except using 1 ng of genomic DNA.

Field trial establishment

The field trial was located at a lawn on the Macdonald Campus, McGill University

(Ste-Anne de Bellevue, QC, Canada). The soil was loamy sand (coarse sand 9%, fine

sand 82%, silt 5%, and clay 4%) with 6.3% organic matter at pH 6.6. The turf was

infested with dandelion and white clover. The S. minor IMI 344141 (isolate 13)

bioherbicide, formulated in barley grain, was applied at 40 g m�2 in June 2006 onto

four different plots. Soil samples were collected from 2 to 7 cm depth in a 25 cm2 area

near the center of the treated plots 5, 15, 30, 60, and 90 days after application. Soil

samples were also collected just prior to application and 60 days after application

from untreated control plots. In addition, plant tissues including dandelion leaves

and turf grass leaves were collected 5 and 15 days after application. Three

replications were collected at each sampling time.

Another turfed field on the Macdonald Campus of McGill University was set to

investigate long-term survival of IMI 344141 in a natural environment. A

randomized complete block design with three replications was set in the field. The

selected field area was on a loamy sand soil (coarse sand 9%, fine sand 80%, silt 5%,

and clay 6%) with 7% organic matter at pH 6.7. The field was separated into three

different treatments consisting of treatment with granular formulation of IMI

344141 at 40 g m�2, treatment with IMI 344141 at 40 g m�2 combined with grass

over-seeding at 15g m�2 and control. The field was treated twice in May and

September 2004. Soil samples were collected in each plot from 2 to 7 cm deep with a

soil sample core (2 cm diameter), and just before application and 9 months after the

second application.

Soil samples were mixed thoroughly to insure uniform distribution and sieved

through a 1-mm diameter sieve to remove any plant materials and stones before

being placed in 50 mL conical tubes. The soil was then flash frozen and kept

at�808C until DNA extraction. DNA extraction was performed separately for

each sample core and PCR was performed twice per extracted sample. Plant

tissues were also frozen in liquid nitrogen and kept at�808C prior to DNA

extraction.

946 P. Li et al.


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Results

RAPD analysis of Sclerotinia minor strains and heterogeneous species

Among all arbitrary 10-mer primers attempted in the RAPD analysis, 35 primers

produced polymorphic DNA band patterns in the first round of screening. In the

second round of screening using DNA of all S. minor isolates and heterogeneous

organisms, OPJ07 (CCTCTCGACA), OPBA14 (TCGGGAGTGG), OPQ13

(GGAGTGGACA) and OPAA15 (ACGGAAGCCC) generated polymorphic band-

ing patterns that were consistently reproducible. However, none of the primers

amplified a unique fragment of IMI 344141 distinct from other isolates.

Among those RAPD primers, OPJ07 (Figure 1) and OPBA14 (Figure 2) were

the most promising candidates. OPJ07 consistently produced a unique band from

Figure 1. OPJ07 (CCTCTCGACA) RAPD profiles of DNA from (1a)(1b) Sclerotinia minor

isolates. (1c)(1d) heterogeneous organisms (fungi, plants and bacteria). Refer to Table 1 and Table

2 for details on the original designation. M�100 bp DNA ladder. N�PCR negative control

(sterile distilled water). Electrophoresis number 13 is the bioherbicide strain IMI 344141 of

Sclerotinia minor. Arrow markers indicate the 737-bp fragment that highlights strain IMI 344141.



15A

isolate 13 (737 bp), but a later PCR result using a specific primers developed from the

sequence of the amplicon737 revealed that the sequence of the amplicon737 was

identical with five other S. minor isolates (isolate 7, 10, 15, 22 and 28) although it

never appeared in these five isolates in RAPD profile, possibly due to insensitivity of

the annealing area of DNA to the primer. OPBA14 consistently produced a

reproducible unique band (1423 bp) from isolate 13, 2 and 5.

Development of SCAR markers for diagnostic PCR

Nucleotide sequencing confirmed the sizes of DNA fragments with the RAPD

primer sequences at both ends of OPJ07 (737 bp, Seq. 1) and OPBA14 (1423 bp, Seq.

2). The sequences were compared to nucleotide sequences of other organisms using

BLASTN. Amplicon737 of OPJ07 had no significant match with sequences of other

organisms. Amplicon1423 of OPBA14 partially hit the sequence of Sclerotinia

Figure 2. OPBA14 (TCGGGAGTGG) RAPD profiles of DNA from (2a)(2b)(2c) Sclerotinia

minor isolates and heterogeneous organisms (fungi, plants and bacteria). Refer to Tables 1 and

2 for details on the original designation. M�Lambda DNA/Hind III Plus ladder. N�PCR

negative control (sterile distilled water). Electrophoresis number 13 is the bioherbicide strain

IMI 344141 of Sclerotinia minor. Arrow markers indicate the 1423-bp fragment that highlights

isolate 2, 5 and 13.

948 P. Li et al.


15A

sclerotiorum 1980 hypothetical protein (SS1G_03032) with 90 bp being identical (150

bits, e-value�1e-32).

OPJ07CCTCTCGACAACCGACTAAAAJ7L6ATAACACACAAGTTGATAATAAA

ATCCTCTTCTAATGCATTAATTACACCATCATTATACAGCTTGTGTATATC

ACTAGTCAAACTAGGTGATATCAATTCATCCAGAACCTTCTTATAACCAAACCTAACAGGGATAAAGAATGCACAAGTACCTTTATTTTTCTTACTCTTA

AACTCAAAATAAGAAATAGAAGGATCTATCACAAGGTGTTTGAAATAGT

GAGACTTATTTAGCTTTTCTAAAATACGTATATCACCAAACTCAGTAAAC

TTCGGCTCAATACCACCCCCCTTCCCTTCGGAAGAAAAAGCTCTAATAT

TTTTATTAGCAATATTTATACGCATTGCTAGCGTTGTAGTTGCJF2ATTTGT

TGTAATAATTGTTTTCATAGATTTCATCAAATATTTGGTAGGGTTTAAGC

CAACCTATACATGCATCACTAACCTTTTCCTTAAAATGACCATGCAGGAC

CCAACGTCACTTGGCTTCTCCGTAGGGATTTTGCTGCCGTTCCCCAACG

ACATCCAACAGCTCTCCCATATGTATCACACTACCTCAGTTCGCCTTTCA

CTATTTCTGGACTGCTTATCCTAAAGGGCTCCTCTCCCTTCCACAGCTTC

ATTJR2TCGACTCGCTTACCTTATGCAACCCJ7R1ATATAGTTTTCGAAGGG

ACTTAGGCCTCGCAGGGTAGCATTCACATTATTTGTTTTCCTGTCGAGAG

GOPJ07

Seq. 1. Amplicon737 produced by RAPD primer OPJ07 (737 bp)

OPJ07 are in bold characters; other primers are underlined.

OPBA14TCGGGTGGCCTGGCTGGTCGTAGCGBA52ATGBA53TCGCTAGGCTGTG

GTGTCTTGGAAGACACGGTGTCGTTACTTGTCGAGTCGGTACTTGAAG

CAGTAATGGTGCTCAAAGTGTTACGAGGTTTGCCGTCTAGATCACGTGG

AAGTGCTCTAGGCTCTCGAGGTAGGTTTATAACTAGGCGTCAAGCAGG

GACTTAAAGTAATGACAATAATATGATGCAATTCGTAACTAATAGTATG

GTATACTAAATGATTGATCTTGAGCACGAAGCTCTACAAAGGGAATCCA

TCCTCCTTATATAGACCTTCATGCTTAGAGGCGCGAAGGGAGCGAGAG

GATCGAGCTTCGATCTCTTCACTCTCCTCGCGCGAAGGGCTCGAGAA

GCCAAGCCTTCGAGCTCTTCGCTCCTTAGCATCACGTGTATGGTGATCC

CGTGACCTAATTGTTACCGGTCGTAACATACAAAGGTATACCGAAGGTT

TATCGACAACAAGGTTATTGCTAGTAATTACACGGAACCTGCCGGCGTC

ATGCTGAAGTGATTGTGACAATAATCATTTAATTATTATTAGCTTCTGAT

TTTTTATTTATTCATTCGTTTATTCATTATTCCATCAATCTCATTCCTTTCATTCATAGAGGTTATCCATATACCAATGCATCCATAACGCCCATCTTATCC

TCCGAAATTCAATACTATACTCCCCATCTGTGTTGATCCATTGAAAGTCC

CAAGTATCTTCTCCTGCACATCCAGCAGATACATCAATCTCTAGTAAACC

ATTCATCACCATCACCATCACCATCCCCACCACCCATTCATCTCTCCTCA

CAACCCATCTATGTACCACCAGTATACAATCCAATTCACATATCCATCTA

GCCAGTCTAGCCCAGCCTAACCTTCCAAACATCCACAATAATGCACAAT

ATACACCTCAAACTTCCTTCCTGGTATTAACGTTCCCACCCACAAATTCC

TCCTCCCACTCAAATTTCGGTAAAATAAGCGATTGACGATCACCCCTTC

TTTTTTCTGCATCTATTTACTGTTAATTGGTTATTCATTTGATTATGAGTA

TATATCCAAATTCATATCTGAGCCCATATGCATATTTTTAATAAATATAAA

GCAAAATTCCAGGACAGATAGAAAGAAAGAAATAAACAATGAAATGGA

AATGAGACTCAAGTAAGTTTATACATACCAATATCAATTTTATCCCCTCC



15A

CATACCAATACCCATACCCATTCCATCACCCGCATATCCACCTACACCTC

TCAGAAACTCAGCCGGTATGGTATTAGAATTAGAATCGACTTTTTCAGC

ATGTACGCCCCATCCTTGCGTATGCATACCCTCTACACCTCCACTCGAAC

TCCTCAAATACCCAABA33TTCCCATCCCCBA31ATCCCCATTCCCACTCCCGAOPBA14

Seq. 2. Amplicon1423 produced by RAPD primer OPBA14 (1423 bp). OPBA14 is in

Italic; BA52 and BA31 are underlined; BA53 and BA33 are in bold characters.

To discriminate the bioherbicide strain (isolate 13) from other isolates, several sets of

SCAR primers were designed from the amplicon737 of OPJ07 and the amplicon1423of OPBA14. When used alone, OPJ7-L6/R1 (Figure 3) highlighted a 669-bpfragment from six isolates of S. minor (isolates 7, 10, 13, 15, 22 and 28) which

were shown to be identical from sequence analysis. BA14-52/31 (Figure 4)

highlighted a 1414-bp fragment from three isolates of S. minor (isolates 2, 5 and

13). All three isolates were fully sequenced and confirmed that they shared 99%

similarity in their sequences. Therefore by combining the results from OPJ07 (OPJ7-

L6/R1) and OPBA14 (BA14-52/31) primers, isolate 13 can be discriminated from the

other 53 samples included in this study.

In order to increase the sensitivity of PCR detection, nested PCR primers weredesigned from the amplicon669 produced by OPJ7-L6/R1 and from the ampli-

con1414 produced by BA14-52/31. JF2/JR2 (Figure 5) highlighted a 280-bp

fragment from seven isolates of S. minor (isolates 7, 10, 13, 15, 22, 28 and 30).

Amplification did not occur in the remaining samples in this study. BA53/33

(Figure 6) produced a 1392-bp fragment from three isolates of S. minor (isolate 2, 5

and 13). Amplification did not occur in the remaining samples in this study. With

combined nested primers JF2/JR2 and BA53/33, isolate 13 can be discriminated

from all samples included in this study.

SCAR markers detection threshold

The sensitivity of the PCR using both OPJ07 and OPBA14 SCAR primers wasexamined on genomic DNA of isolate 13. To determine the threshold detection

limit of a DNA concentration at which PCR with the primers can produce a

visible product in gel electrophoresis, genomic DNA of isolate 13 was serial

diluted from 100 pg to 1 fg per 25 mL reaction solution. The lowest concentration

of genomic DNA at which a visible band was produced was 10 pg per reaction

when PCR was performed with OPJ7-L6/R1 and BA14-52/31 primers at first

round PCR. The sensitivity was increased 1000 times to 10 fg when the first-

round PCR product was amplified again with nested primer JF2/R2 and BA53/33, respectively.

Detection of sclerotia

As with most soilborne diseases, sclerotia of S. minor are mostly clumped or

clustered in field soil. Although it was observed that S. minor IMI 344141 applied in

the field rarely produced sclerotia and they did not survive over winter to the degree

to reproduce the disease to plants in the following season in nature (PMRA 2007;

950 P. Li et al.


15A

Watson 2007), it is essential to know if sclerotia can be detected in soil using our

molecular markers, and what is the minimum quantity of sclerotia needed for

detection.

A sclerotium was ground in 0.2 g of soil and 1 ng of the extracted genomic DNA

was used for PCR. With ITS-1F/ITS4 primers, fungal DNAwas shown to be present

in all soil samples. In first round PCR, a weak band appeared with J7L6/R1 and no

band for BA14-52/31. In nested PCR, the band was intense from both JF2/R2 and

BA53/33. This demonstrated that one single sclerotium of S. minor IMI 344141 can

be detected from soil and PCR amplification showed little difference between

autoclaved soil and natural soil.

Figure 3. Amplicon669 produced by OPJ7-L6/R1. (3a)(3b) DNA from Sclerotinia minor

isolates. (3c)(3d) DNA from heterogeneous organisms (fungi, plants and bacteria). Refer to

Tables 1 and 2 for details on the original designation. M�1 kb Plus DNA ladder. N�PCR

negative control (sterile distilled water). Electrophoresis number 13 is the bioherbicide strain

IMI 344141 of Sclerotinia minor. Arrow markers indicate the 669-bp fragment that highlights

strain IMI 344141 and other 5 isolates of S. minor (isolate 7, 10, 15, 22 and 28).



15A

Persistence of Sclerotinia minor IMI 344141

Both markers were detected during first round PCR (BA14-52/31 and OPJ7-L6/R1)

and nested PCR (BA14-53/33 and JF2/R2) from infected plant tissues which were

collected 5 and 15 days after S. minor application in the field (Figures 7 and 8). IMI

344141 (isolate 13), formulated in barley grits, was applied in field trials at different

times (Figure 8). By combined OPJ7-L6/R1 and BA14-52/31 for the first round PCR,

JF2/R2 and BA53/33 for nested PCR, S. minor can be successfully tracked in soil.

The marker fragments did not appear in soil samples that were collected from the

control plots before application, 2 and 9 months after application, indicating that

S. minor was not present in the control field. From first round OPJ7-L6/R1 screening

with all other soil samples (Figure 8a), good bands appeared from 5- and 15-day

samples, very weak bands from 1-month samples, and no band from samples taken

on or after 2 months. From nested PCR using JF2/R2 (Figure 8b), 5-day, 15-day,

1-month and 2-month samples had intense bands, but no marker DNA was

amplified from samples taken 3 and 9 months after bioherbicide application.

Figure 4. Amplicon1414 produced by BA14-52/31 (4a)(4b) DNA from Sclerotinia minor

isolates and heterogeneous organisms (fungi, plants and bacteria). Refer to Tables 1 and 2 for

details on the original designation. M1�Lambda DNA/Hind III Plus ladder. M2�100 bp

DNA ladder. L�Romaine lettuce. ITS�Fragment (�530 bp) produced from mycelia DNA

of isolate 13 with ITS-1F/ITS4 primers. N�PCR negative control (sterile distilled water).

Electrophoresis number 13 is the bioherbicide strain IMI 344141 of Sclerotinia minor. Arrow

markers indicate the 1414-bp fragment that highlights isolate 2, 5 and 13.

952 P. Li et al.


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From first round BA14-52/31 screening with all other soil samples (Figure 8c,d),

no band amplified from the soil samples. From nested PCR using BA53/33 (Figure

8e), 5-day, 15-day, 1-month and 2-month samples had intense bands, but no marker

DNA was amplified from samples taken 3 and 9 months after application. Thus inthis 9-month field trial, two-round PCR using both markers demonstrated that

S. minor isolate IMI 344141 was still present in soil 2 months after application, but

did not appear in 3- and 9-month soil samples.

Discussion

It is imperative when applying a biological control agent in the field that the strain

released remains within the area of application (Zhou, Bailey and Derby 2004). This

is sometimes difficult to ascertain with traditional techniques, especially when the

agent is a fungus which rarely forms characteristic sexual structures. Apothecia andascospore production in S. minor is very rare in the field (Hind, Ash and Murray

2001) and has not been recorded to occur in North America (Subbarao 1998).

Dispersal and transmission of the disease is exclusively by the direct contact with

germinating sclerotia to produce infective hyphae which colonize plants and

Figure 5. Amplicon280 produced by JF2/R2 (5a)(5b) DNA from Sclerotinia minor isolates

and heterogeneous organisms (fungi, plants and bacteria). Refer to Tables 1 and 2 for details

on the original designation. M1�Lambda DNA/Hind III Plus ladder. M2�100 bp DNA

ladder. L�Romaine lettuce. ITS�Fragment (�530 bp) produced from mycelia DNA of

isolate 13 with ITS-1F/ITS4 primers. N�PCR negative control (sterile distilled water).

Electrophoresis number 13 is the bioherbicide strain IMI 344141 of Sclerotinia minor. Arrow

markers indicate the 280-bp fragment that highlights isolate 7, 10, 13, 15, 22, 28 and 30.



15A

eventually produce more sclerotia which are returned to the soil. When applied as a

bioherbicide to turfgrass, sclerotia rarely formed and the eruptive mycelium of the

S. minor bioherbicide does not persist in the absence of a susceptible host, and

quickly decays within 10 days (Watson 2007). When applied as an integrated

microbial pest control product (MPCP) onto turfgrass S. minor IMI 344141 rarely

produces sclerotia and apothecia have never been observed. Sclerotia or infective

hyphae of S. minor IMI 344141 does not persist in the turf environment.

In this study, RAPD-PCR was used to develop markers that could be used to

differentiate isolates of S. minor from a worldwide collection of the fungus. Many

primers were able to be used to differentiate isolates of the fungus, but a combination

of two primers had to be used to clearly differentiate the bioherbicidal strain IMI

344141 from all other isolates of S. minor. The specificity and the sensitivity of the

markers were improved by the development of SCAR markers based on the RAPD

primers and the use of nested PCR. These markers could be used to detect a single

Figure 6. Amplicon1392 produced by BA14-53/33 (6a)(6b)(6c) DNA from Sclerotinia minor

isolates and heterogeneous organisms (fungi, plants and bacteria). Refer to Tables 1 and 2 for

details on the original designation. M�100 bp DNA ladder. L�Romaine lettuce.

ITS�Fragment (�530 bp) produced from mycelia DNA of isolate 13 with ITS-1F/ITS4

primers. N�PCR negative control (sterile distilled water). Electrophoresis number 13 is the

bioherbicide strain IMI 344141 of Sclerotinia minor. Arrow markers indicate the 1392-bp

fragment that highlights isolate 2, 5 and 13.

954 P. Li et al.


15A

sclerotium in soil and had a detection limit of 10 fg of S. minor DNA. Detection of

the fungus from soil is dependent upon a rigorous DNA extraction protocol and

testing of the proposed primers against a wide range of sources of DNA which are

likely to be encounted within the field situation. This is especially important when

dealing with pathogens with potentially wide host ranges (Ash 2010). By applying

these techniques to a field trial where the fungus had been applied, the S. minor could

be detected for up to 3 months post application. It is proposed that this DNA is from

living cells as it has been shown that naked DNA within the soil is quickly broken

down in 2�5 days (Romanowski, Lorenz, Sayler and Wackernagel 1992). The

protocol also showed that the fungus could not be detected after 9 months post

application. As this strain of the fungus has not been observed producing sclerotia in

field situations, it does not colonize grass and mycelia of the fungus do not survive

beyond 11 days in the turfgrass environment, the likelihood of the fungus spreading

from, or persisting in the area of application is improbable. Furthermore, unlike the

studies of Zhou et al. (2004; Zhou, Bailey, Chen and Keri 2005) with P. macrostoma,

the S. minor is applied as a colonized grain formulation, and so drift from the site of

application is highly unlikely.

Ideally, a single pair of PCR primers could be designed to allow the identification

and tracking of the strain of biocontrol agent used. However, the use of a

combination of SCAR markers as reported here is an effective means of

differentiating strains of a largely clonal fungus such as S. minor. In the future, the

primers could be combined in a single multiplex PCR allowing the assay to be

undertaken in a single tube. This approach has been used to differentiate species of

Figure 7. Detection of Sclerotinia minor IMI 344141 in plant tissue in the field (7a)

amplification from J7L6/R1, (7b) amplification from BA14-52/31. M�100 bp DNA ladder.

1, 2, 3, 7, 8, 9: plant tissues collected 5 days after application of S. minor IMI 344141;

4, 5, 6, 10, 11, 12: plant tissues collected 15 days after application of S. minor IMI 344141;

N1�nested PCR from 1; N2�nested PCR from 2, etc. P�positive control amplified with

mycelia DNA of isolate13 using BA14-52/31; N�negative control with sterile distilled water

using BA14-52/31; NP�positive control amplified with mycelia DNA of isolate13 using

BA14-53/33; NN�negative control with sterile distilled water using BA53/33; JP�positive

control amplified with mycelia DNA of isolate13 using OPJ7-L6/R1; JN�negative control

with sterile distilled water using OPJ7-L6/R1; NJP�positive control amplified with mycelia

DNA of isolate13 using JF2/R2; NJN�negative control with sterile distilled water using

JF2/R2.



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Figure 8. Field trial of isolate IMI 344141 of Sclerotinia minor with OPJ07 and OPBA14

SCAR markers: (8a) first round using OPJ7-L6/R1. (8b) nested PCR using JF2/R2. (8c) (8d)

956 P. Li et al.


15A

Sclerotiniaceae using the lacc2 gene by Hirschhauser and Frohlich (2007). The

inclusion of specific primers for a gene such as the lacc2 (Hirschhauser and Frohlich

2007) or non-coding region such as the ITS (White et al. 1990) would reduce the risk

of obtaining a false-negative result (O’Brien 2008). This current research could also

be used as a basis to develop a quantitative PCR assay to more accurately define the

environmental fate of Strain IMI 344141 of S. minor in the field as demonstrated by

Dauch et al. (2006) using C. coccodes.

In summary, this research has clearly shown that SCAR markers developed from

RAPD profiles of S. minor can be optimized to differentiate the bioherbicidal strain

of S. minor (strain IMI 344141) from other strains of the fungus and DNA from

other organisms commonly found in turfgrass soil. Furthermore, the sensitivity of

the assay developed allows the tracking of the organism in the field and adds

evidence that S. minor (strain IMI 344141) degrades rapidly in the turfgrass

environment and as such is expected to have minimal impact on the environment.

Acknowledgements

Financial support from the Natural Sciences and Engineering Research Council of Canada(NSERC) Idea to Innovation (I2I) grant and 4260864 Canada Inc (Sarritor Inc.) are gratefullyacknowledged. The following graciously loaned S. minor isolates: G. Abawi, CornellUniversity, Geneva, USA; G. Boland, University of Guelph; J. Whipps, Warwick HorticultureResearch International, UK; H. Huang, Agriculture and Agri-food Canada, Brandon;P. Phipps, Virginia Tech; S. O’Neill, Queensland Department Plant Industry, Brisbane;Tamrika Hind-Lanoiselet, Wagga Wagga Agricultural Institute, NSW; S. Morley, HerbariumVPRI, Victorian Department of Primary Industries, Knoxfield, Victoria, Australia; K. Elena,Benaki Phytopathological Institute, Athens, Greece; Alison Stewart, Lincoln University,Christchurch, New Zealand; Levente Kiss, Hungarian Plant Protection Institute, Budapest; P.Charue, Belgian Coordinated Collections of Microorganisms, Louvain; J. Tatnell and M.Fuhlbohm, Australian Department of Plant Industry, Kingaroy, QLD; W.G. Kim, NationalInstitute of Agricultural Science and Technology, Suwon, Republic of Korea; F. Snippe-Claus,Centraalbureau voor Schimmelcultures, Utrecht, The Netherlands.

first round using BA14-52/31. (8e) nested PCR using BA14-53/33.

M�Lambda DNA/Hind III Plus ladder.

C01C02C03� three samples collected from control plot before application; S11S12S13�5

days after application; S21S22S23�15 days after application; S31S32S33�1 month after

application; 2M12M22M3�2 months after application; C21C22C23�soil samples from

control plot after 2-month application; 3M13M23M3�3 months after application;

C91C92C93�soil collected from control plot for 9-month trial before application;

S919293�soil samples collected from control plot after application for 9 months;

S949596�9 months after application; S979899�9 months after application, treatment

combined with grass over-seeding;

P: positive control using mycelia DNA of isolate 13 with OPJ7-L6/R1; N: negative control

replaced DNAwith sterile distilled water using primer OPJ7-L6/R1; NP: positive control using

mycelia DNA of isolate 13 with JF2/R2; NN: negative control using JF2/R2; NJP: positive

control using primer JF2/R2 with the DNA template from first round PCR product of P

(OPJ7-L6/R1); BP: positive control using mycelia DNA of isolate 13 with BA14-52/31; BN:

negative control replaced DNA with sterile distilled water using primer BA14-52/31; NBP:

positive control using mycelia DNA of isolate 13 with BA14-53/33; NBN: negative control

using BA14-53/33.



15A

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RESEARCH ARTICLE

Physiological characterization of the dandelion bioherbicide, Sclerotiniaminor IMI 344141

In’aam Y. Shaheena, Mohammed H. Abu-Dieyeha$, Gavin J. Ashb and

Alan K. Watsona*

aDepartment of Plant Science, McGill University, 21,111 Lakeshore Road, Ste-Anne-de-Bellevue, Quebec, H9X 3V9 Canada; bE.H. Graham Centre for Agricultural Innovation (NSWDepartment of Primary Industries and Charles Sturt University), Charles Sturt University,

Locked Bag 588, Wagga Wagga, NSW, 2678, Australia

(Received 26 February 2009; returned 6 April 2009; accepted 15 October 2009)

The fungus Sclerotinia minor (IMI 344141) is being developed as a biologicalcontrol for dandelion and other broadleaf weeds in turfgrass environments. Beinga microbial pest control agent (MPCA), the S. minor strain must be characterizedto show relatedness to like organisms and to distinguish the MPCA from relatedmicroorganisms. Phenotypic variation among 30 isolates of S. minor, collectedfrom different regions and hosts, was studied on potato dextrose agar (PDA)and oatmeal agar (OMA). Isolates varied significantly in sclerotia shape (length/width ratio) and number, but did not vary in colony morphology or growth rates.There was high diversity (0.6) among the mycelial compatibility groups (MCG) asseven multi-member and 11 single-member groups were recognized. Isolates werecategorized into highly virulent, virulent, moderately virulent, and hypo virulentbased on 48 h post mycelial growth on detached dandelion leaves. When assessedon dandelion plants in the greenhouse, isolate IMI 344141 ranked the highest inbiocontrol efficacy, reduction of above- and below-ground biomass, andreduction in dandelion survival. Oxalic acid production was not correlated withisolate aggressiveness or growth rate and did not vary among isolates of the sameMCG. IMI 344141 can be phenotypically distinguished from the other tested S.minor isolates by performing vegetative compatibility testing and countingsclerotia produced on standard 9-cm diameter PDA plates. IMI 344141 producesB100 sclerotia/plate.

Keywords: phenotypic variation; mycelial compatibility; virulence; oxalic acid

Introduction

Plant pathogenic fungi regularly cause significant crop losses. However, many weed

species are also attacked by fungi which have led to investigations of fungi as

biological weed control agents. One such fungus, Sclerotinia minor Jagger, is

pathogenic to many plant species, and several studies have investigated the

pathogenicity of S. minor on dandelion (Taraxacum officinale Weber ex Wiggers)

and other broadleaf weeds in turfgrass (Ciotola, Wymore, and Watson 1991; Riddle,

*Corresponding author: Email: [email protected]$Present address: Department of Biology and Biotechnology, The Hashemite University,

PO Box 150459, Zarqa, Jordan.

ISSN 0958-3157 print/ISSN 1360-0478 online

# 2010 Taylor & Francis

DOI: 10.1080/09583150903419520

http://www.informaworld.com

Biocontrol Science and Technology,

Vol. 20, No. 1, 2010, 57�76

Downloaded By: [Canadian Research Knowledge Network] At: 14:53 12 January 2010

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Burpee, and Boland 1991; Briere, Watson, and Paulitz 1992; Schnick, Stewart-Wade,

and Boland 2002; Stewart-Wade et al. 2002; Abu-Dieyeh, Bernier, and Watson 2005;

Abu-Dieyeh and Watson 2006, 2007a,b,c). The IMI 344141 strain of S. minor is

registered in Canada as a bioherbicide for control of dandelion in turfgrass

environments (PMRA 2007).

S. minor is a soil-borne Discomycete (Sclerotineaceae) fungus characterized bysmall (0.5�2.00 mm) spherical sclerotia (Willetts and Wong 1980) that germinate by

eruptive growth of the mycelium and colonize susceptible plant tissues (Abawi and

Grogan 1979). The host range of S. minor is broad and includes 21 families, 66

genera and 94 species (Melzer, Smith, and Boland 1997; Hollowell, Shew, Cubeta,

and Wilcut 2003). S. minor is a causal agent of lettuce drop, Sclerotinia blight of

peanut, white mold of green beans, and watery soft rots of vegetables (Abawi and

Grogan 1979). In Canada, S. minor is not considered as a serious plant pathogen

except on lettuce (Melzer et al. 1997).

Individual strains of a broad spectrum fungus may vary in their ability to infect

different hosts. Therefore, the development of bioherbicide formulations based on

selected isolates requires the capability to distinguish the biocontrol agent from like

organisms (Noonan, Glare, Harvey, and Sands 2004). Accurate identification of the

microbial pest control agent (MPCA) is a key component of safety assessment and

taxonomic designation to an appropriate level to distinguish the MPCA from related

microorganisms is a regulatory requirement (PMRA 2001). The ability to

discriminate the IMI 344141 bioherbicide strain from other S. minor isolates is

also imperative for environment tracking and intellectual property and litigationprotection.

Mycelial compatibility has been used to assess the population biology of many

fungal plant pathogens. Mycelial compatibility is the ability of two strains of fungi to

fuse and form a stable heterokaryon (Leslie 1993). Strains that are compatible with

one another are frequently described as members of the same mycelial compatibility

group (MCG). Mycelial compatibility reflects genetic heterogeneity among isolates

(Viji, Uddin, O’Neill, Mischke, and Saunders 2005), reveals intrapopulation changes

(Sarma and Singh 2002), measures and categorizes intraspecific variations (Kohn,

Carbone, and Anderson 1990; Leslie 1993, 1996; Durman, Menendez, and Godeas

2003), and if combined with molecular markers, gives more precise information

about genetic diversity and relatedness among isolates of the same species (Kohn,

Stasovski, Carbone, Royer, and Anderson 1991; Noonan et al. 2004).

Pathogenic strains that are vegetatively compatible are presumed to originate

from the same clone, even if they are geographically isolated from one another

(Leslie 1993). Correlation between MCGs and other characters such as pathogeni-

city or phenotype could lead to useful diagnostics. Widely dispersed MCGs ofS. sclerotiorum showed varied aggressiveness while isolates from the same field did

not vary in aggressiveness (Kull and Pedersen 2004). Isolates of Sclerotium rolfsii

from one MCG varied in the number and size of sclerotia in culture, indicating that

phenotypic differences are not uncommon among members of a MCG (Punja and

Grogan 1983). In other studies, morphological or pathological characters were not

related to mycelial compatibility grouping (Durman et al. 2003).

The physiology of pathogenesis of plant diseases caused by Sclerotinia species has

been comprehensively reviewed and this has clarified the importance of cell wall-

degrading enzymes and oxalic acid (OA) production by the pathogen in causing

58 I.Y. Shaheen et al.


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disease (Lumsden 1979; Willetts and Wong 1980). However, pathogenesis of S. minor

specifically is not well understood. With other pathogenic Sclerotinia species, OA is

implicated in the infection process (Bateman and Beer 1965; Kritzman, Chet, and

Henis 1977), but contradicting correlations have been demonstrated between OA

production and virulence. Oxalic acid has been confirmed as a pathogenicity

determinant in S. sclerotiorum by using an OA-deficient mutant (Godoy, Steadman,

Dickman, and Daur 1990), but OA was not the sole pathogenic determinant of S.

trifoliorum (Callahan and Rowe 1991). Variation in the production of OA by isolates

of some Sclerotinia species has been reported, while few studies have examined OA

production by S. minor. One study reported correlation between aggressiveness of S.

minor isolates on peanut and mycelial growth in broth cultures, but not OA

production (Hollowell, Smith, and Shew 2001).

This study was designed to demonstrate the uniqueness of the IMI 344141

bioherbicide isolate of S. minor by comparison to other S. minor using phenological

traits, MCGs, aggressiveness, and OA production. Experiments were conducted to:

(1) detect morphological variability among S. minor isolates collected from different

regions in the world; (2) examine relatedness amongst the isolates based on mycelial

compatibility interactions; (3) compare the pathogenicity of the different isolates and

their affinity to produce oxalic acid; and (4) provide a set of descriptors to

distinguish the IMI 344141 isolate of S. minor from related microorganisms.


Fungal Isolates

Thirty isolates of S. minor were obtained from various culture collections and

research colleagues (Table 1). The isolates were maintained by growing single

sclerotia, or agar discs taken from the original cultures, and growing them on potato

dextrose agar (PDA) (Difco Laboratories Inc, Detroit, MI) at 20918C in the dark.

For long-term maintenance, two sets of PDA slants were inoculated with each

isolate; one set was placed under sterile mineral oil and the other under �80928C.Sclerotia of each of the 30 isolates were produced on surface sterilized market

dandelion leaves in moist chambers. Leaves were inoculated with mycelial discs taken

from the margin of 4�5-day-old colonies grown on PDA and incubated at 20918C in

the dark. Collected sclerotia were air-dried at 20918C for 48 h, labeled, and stored in

air-tight glass vials at 490.58C.

Experiment 1. Morphological variation

Cultures of each of the 30 S. minor isolates were established by sterilizing sclerotia

(prepared above) in 70% ethanol for 40 s, 5% bleach solution for 3 min, washing

twice with sterilized distilled water, and drying on sterilized filter paper (Abu-Dieyeh

and Watson 2006). Surface sterilized sclerotia were transferred to the surface of PDA

in 9-cm diameter Petri plates and incubated at 20918C in the dark. Three-millimeter

diameter mycelial discs were taken from the margin of 4�5-day-old colonies grown

and used to inoculate PDA and oatmeal agar (OMA) (Sigma-Aldrich Canada Ltd,

Oakville, ON) plates. Plates were incubated at 20918C in the dark and examined

daily for colony diameter, mycelial density and color, sclerotial initiation and



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formation times, sclerotial arrangement, exudate presence on sclerotia, and sclerotial

size. Colony diameter was measured daily until the growth reached the edge of the 9-

cm diameter Petri plates. The experiment was replicated three times. After 20�25days of growth, the total number of sclerotia on PDA plates was counted and the

diameter of 25 sclerotia formed on both growth media per isolate per replicate was

measured; the data from the replicated plates were averaged (Sarma and Singh 2002).

Experiment 2. Mycelial compatibility groups

Each isolate was paired against itself as a control and against all other isolates on

modified Patterson’s medium (MPM) (Kohn et al. 1990). This medium contains

0.68 g KH2PO4, 0.5 g MgSO4.7H2O, 0.15 g KCl, 0.5 g yeast extract, 1 g NH4NO3,

Table 1. Designation codes, origin, host plant and mycelial compatibility group (MCG) of a

worldwide collection of Sclerotinia minor isolates.

MCG Isolate code/number Origin Host

1 IMI 344141 Quebec, Canada Lettuce (Lactuca sativa)

LRC2104 Manitoba, Canada Lettuce (Lactuca sativa)

ATCC44236 New York, USA Lettuce (Lactuca sativa)

Sm44 California, USA Lettuce (Lactuca sativa)

Sm66 Ontario, Canada Lettuce (Lactuca sativa)

2 S96-22 Suwon, Korea Unknown

S96-138 Suwon, Korea Unknown



3 R21 Virginia, USA Peanut (Arachis hypogaea)

R23 Virginia, USA Peanut (Arachis hypogaea)

4 R22 New Jersey, USA Lettuce (Lactuca sativa)

R25 New Jersey, USA Cabagge (Brassica campestris)

5 BRIP28139 New South Wales, AU Chickpea Cicer arientium

K4205 Queensland, AU Mungbean (Vigna radiata)

6 VPRI1671 Tasmania, AU Onion (Allium sativum)

VPRI1285 Victoria, AU Bathurst burr (Xanthium spinosum)

7 BPIC1681 Kifissia, Greece Lettuce (Lactuca sativa)

BPIC1949 Kifissia, Greece Lettuce (Lactuca sativa)

8 R24 New Jersey, USA Endive (Cichorium endivia)

9 JW1 Hull, UK Unknown

10 SMRF02 Virginia, USA Peanut (Arachis hypogaea)

11 TH1C New South Wales, AU Canola (Brassica napus)

12 LU286 Canterbury, NZ Lettuce (Lactuca sativa)

13 LK1 Budapest, HU Knapweed Centaurea pannonica

14 MUCL38484 Belgium Unknown

15 S94-001 Suwon, Korea Unknown

16 CBS112.17 The Netherlands Lettuce (Lactuca sativa)

17 CBS207.25 The Netherlands Sasify (Tragopogon porrifolius)

18 CBS339.39 Italy Lettuce (Lactuca sativa)



15A

18.4 g d-glucose, 0.2 mL of Vogel’s trace elements solution, 15 g agar (Fisher

Scientific Company, Ottawa, ON) and six drops of McCormick’s food coloring.

Each isolate was grown on MPM for 5 days prior to pairing. For pairings, mycelial

discs (3-mm diameter) taken from the edge of an actively growing colony of eachisolate were placed approximately 30 mm apart on MPM in 9-cm diameter Petri

dishes, one pairing per dish, and incubated at 20918C in the dark. Isolates were

paired in all combinations; each pairing was performed three times (three replicates).

Pairings were examined visually 7�15 days post inoculation for the presence of an

antagonistic zone. Pairings were assessed as incompatible if there was a red line in the

reaction zone between the two colonies, and as compatible when the two colonies

merged with no detectable line (Kohn et al. 1990, 1991; Sarma and Singh 2002;

Durman et al. 2003). Questionable pairings were repeated to verify results.

Experiment 3. Oxalic acid production

Cultures of the 30 S. minor isolates were established by surface sterilizing sclerotia of

each isolate as mentioned previously and plating on PDA. Plates were incubated at

20918C in the dark until mycelial growth reached two-thirds of the plate. Five

mycelial discs (6-mm diameter) taken from the margin of an actively growing colony

of each isolate were used to inoculate 100 mL potato dextrose broth (PDB, Difco).

Flasks were placed on a rotary shaker (50 rpm) at 20918C in the dark for 7 days.

Cultures were filtered through P8 filter Whatman paper (Fisher) and each filtrate

was filtered through a series of Millipore membrane filters with decreasing pore size8, 5, 1 and 0.45 mm (Millipore Corporation, Bedford, MA) using a filter holding

system (Nalgene Nunc. International, Rochester, NY) and a vacuum pump. Fifteen-

microlitre aliquots taken from the filtrate of each fungal isolate were injected into an

HPLC system utilizing an Alltech Prevail Organic Acid 5 mm Column (150�4.6 mm

ID) for organic acid analysis (Alltech Associates Inc., Deerfield, IL) and the Waters

HPLC system with a 441 Detector (Millipore Corporation, Milford, MA). The

elution solvent was 25 mM KH2PO4, pH 2.5. Organic acid detection was carried out

at 210 nm at 258C. Ten microlitres of the oxalic acid standard was chromatographedbefore the samples. Concentrations were calculated by comparing peak areas of the

samples with those of oxalic acid standard solutions. The experiment was conducted

in a completely randomized design with three replications and repeated three times.

Experiment 4. Virulence

Virulence of the S. minor isolates was determined on detached dandelion leaves and

on whole plants in the greenhouse. Mycelial discs and colonized barley grits were

used to screen the isolates on detached dandelion leaves, while the colonized barley

grits were also used on whole plants in a greenhouse to compare the virulence of the

10 most virulent isolates from the detached leaves bioassay.

Plant production

Locally collected dandelion seeds were sown onto potting soil [two-third black

pasteurized soil and one-third Pro-mix (Premier Promix, Premier Horticulture Ltee,

Riviere-du-Loup, QC) in 40�30�8 cm trays in the greenhouse (20928Cwith 15 h of



15A

light day1 at photon flux density minimum of 350950 mmol m�2 s�1)]. Two-week-old

seedlings with uniform vigor were individually transplanted into 15-cm diameter pots

containing the mixed potting soil (as above). Plants were grown in the greenhouse

with programmed drip irrigation of 50 mL pot�1 three times a day.

Detached leaf bioassay

Leaves were removed from 8-week-old dandelion plants grown in the greenhouse and

three dandelion leaves were placed in each of 90 glass Petri dishes lined with

Whatman No. 1 filter paper moistened with 4 mL sterile distilled water. PDA plates

of each of the 30 isolates were prepared as above. Agar plugs (2-mm diameter) were

cut from the actively growing margins of cultures of each of the 30 S. minor isolates

and placed with the mycelium-side down in the center of a detached dandelion leaf

(Briere, Watson, and Hallett 2000; Hollowell and Shew 2003). Petri dishes were

sealed with parafilm and incubated at 20928C in the dark for 48 h. Lesion diameters

were measured (lengthwise direction of the leaves) at 24 and 48 h after inoculation.

The experiment was conducted in a completely randomized design with three plate

replicates (a total of nine dandelion leaves) per isolate and conducted twice. The 48 h

post inoculation screening results were used to arbitrarily classify the S. minor

isolates into four categories: highly virulent (mean lesion diameter ]30 mm),

virulent (20�30 mm), moderately virulent (10�20 mm), and hypovirulent or avirulent

(B10 mm).Highly virulent isolates and IMI 344141 were assessed for their aggressiveness by

growing them on sterilized barley grits as described by Abu-Dieyeh and Watson

(2006) and screening them on detached dandelion leaves using the same procedures

used for mycelial discs.

Greenhouse bioassay

When potted dandelion seedlings were 3-weeks-old, 0.8 g of a commercial grass seed

mixture [30% Kentucky bluegrass (Poa pratensis), 40% creeping red fescue (Festuca

rubra L. var. rubra) and 30% turf type perennial ryegrass (Lolium perenne L.),

(C.I.L.† GolfgreenTM, Spectrum Brands Canada Ltd, Brantford, ON)] was scattered

over the surface of each pot. The grass was cut weekly with hedge shears to a height

of 10 cm, commencing 3 weeks after grass sowing. A 15�15�30, N�P2O5�K2Ofertilizer with micronutrients (PlantexTM, Plant Product Co, Brampton, ON) was

applied at 3.5 g L�1 when the dandelions were 5-weeks-old. Eight weeks after

transplanting, plants were labeled for the specific treatment, 16 plants for each

isolate (eight of which were retained as untreated controls), were distributed in a

completely randomized design on the bench, misted with water, and each was

inoculated with 0.2 g of the S. minor isolates granular formulation. Mist was applied

daily over all the pots for 1 week. Plant survival was recorded weekly for 3 weeks.

Plant re-growth (after 100% above ground damage) was recorded as decreasing

percentage damage by estimating the biomass of the new leaves. Three weeks after

the first application, surviving plants received a second application of 0.2 g of their

S. minor isolates granular formulation. Six weeks after the first application, all

dandelion plants were carefully removed from the soil, the roots were thoroughly

washed and dissected above the crown, separating above- and below-soil biomass.



15A

Treated and control plant materials (leaves or roots) were separately bulked for each

of the 10 isolates, placed in paper bags, oven dried at 808C for 72 h and then weighed.

The experiment was repeated.

Statistical data analysis

Bartlett’s test of SAS (SAS Institute Inc., Cary, NC 2002) was used to test the

homogeneity of variances of data from the two replicates of each experiment. In all

experiments, data from the two repeats were homogeneous and data were combined

and analyzed as one experiment. Colony diameter measurements, sclerotia count per

plate, lesion diameter on dandelion leaves and oxalic acid concentration of the

S. minor isolates were analyzed using one-way ANOVA and the mean values of the

isolates were separated using Tukey’s test at P�0.05. Sclerotia count per plate data

of two isolates, IMI 344141 and CBS339.39 were significantly less than other

isolates, but not significantly different from each other, thus they were subjected to a

paired t-test. One-way ANOVA and mean comparisons for all experiments were

performed using SigmaStat 2.03 (1992�1997).

Results

Experiment 1. Morphological variation

The studied S. minor isolates represent a worldwide sampling of S. minor populations

from more than three different hosts including the dominant host, lettuce (Table 1).

Growth characteristics were tested on PDA and OMA media. Each of the isolates

showed similar appearance of mycelial color (mainly white) and type (mainly fluffy)

on both agars. Sclerotia initiation and formation time was more rapid on OMA than

on PDA. Eight isolates required 1 day from initiation to formation on PDA, while

the 13 isolates needed 1 day on OMA. In general, the time for sclerotia completion

was 1�4 days except for isolate CBS339.39 which was the slowest in initiation and

formation, 10 and 4 days, respectively. Three isolates did not form sclerotia, even

after a month of incubation. Sclerotia formation pattern was different on the two

growth media with mostly concentric on OMA and peripheral to scattered on PDA.

Sclerotia varied in color from olive green, dark brown, grey to dominant black.

Sclerotia were spherical to irregular in shape, most with a rough surface.

The sclerotia of IMI 344141 were almost spherical, black, highly rough surfaced,

1.72�1.69 mm in diameter, and scattered over the surface of PDA. Microspores

were not detected with any of the isolates. Exudates were produced on the sclerotia of

most isolates on OMA but rarely on PDA. An earthy soil odor was clearly evident

for five isolates on OMA and for two on PDA.

Except for isolate CBS207.25, the growth rates on a particular medium were

similar among the isolates (Table 2). In general, radial growth was faster for all

isolates on PDA than on OMA, ranging from 5.3 to 17.3 mm and 5.3 to 15.1 mm

after 24 h on PDA and OMA, respectively, and 26.5 to 57 mm and 15.3 to 37.5 mm

after 48 h (Table 2). The growth rate of IMI 344141 was 13.393.22 and 48.894.25

on PDA after 24 and 48 h, respectively.

Sclerotia size was one of the most obvious quantitative characters with significant

differences among isolates in length and width dimensions. Isolate S94-001 had the



15A

smallest sclerotia on PDA (1.44�1 mm) and Sm66 had the smallest on OMA

(1.16�0.95 mm) (Table 3). While the largest sclerotia on both growth media were

CBS339.39, 3�2.4 mm on PDA and 2.6�2.1 mm on OMA, respectively. The very

large sclerotia suggests CBS339.39 had been misidentified and nearer to

Table 2. Growth rate of Sclerotinia minor isolates after 24 and 48 h of incubation on PDA

and OMA media. Average of four replications.

Colony diameter (mm)

PDA OMA

MCG Isolate 24 h 48 h 24 h 48 h

1 IMI344141 13.393.221 48.894.25 11.893.59 32.394.86

LRC2104 8.094.58 37.798.08 10.396.29 27.0916.02

ATCC44236 17.097.81 54.8911.64 12.695.79 33.098.76

Sm44 5.792.51 34.093.61 11.092.16 29.3910.21

Sm66 6.393.06 35.3910.79 11.394.35 36.097.39

2 S96-22 10.094.58 45.097.21 12.194.37 34.595.45

S96-138 5.392.08 32.3919.50 10.194.91 31.597.33

S96-250 13.095.20 53.8912.71 11.693.68 30.098.12

S97-75 10.794.16 45.394.62 14.893.62 34.696.05

3 R21 17.1796.75 48.0916.52 9.092.71 29.3912.23

R23 10.094.00 47.097.81 14.494.61 38.498.26

4 R22 8.6790.58 33.895.25 8.594.04 34.6913.01

R25 7.6794.04 36.396.35 12,092.94 31.099.49

5 BRIP28139 10.095.57 45.3918.77 10.996.46 30.9910.44

K4205 9.095.20 44.3912.42 11.895.85 29.9911.65

6 VPRI1671 9.093.61 39.398.15 13.392.50 28.097.26

VPRI1285 12.794.93 42.7912.74 10.693.86 29.096.78

7 BPIC1681 9.791.53 28.791.53 15.091.41 33.393.40

PBIC1949 16.096.25 44.0912.77 14.394.19 35.394.57

8 R24 16.6792.31 52.093.61 14.194.73 36.699.43

9 JW1 7.091.73 32.291.76 8.592.65 25.196.76

10 SMRF02 16.398.15 55.7913.65 15.194.37 37.598.35

11 TH1C 6.792.52 36.796.51 9.994.09 28.197.22

12 LU286 14.096.08 48.0911.27 13.892.63 34.897.41

13 LK1 7.792.31 33.093.46 7.091.58 24.9912.07

14 MUCL38484 13.295.80 38.0918.19 11.495.41 33.4910.47

15 S94-001 17.398.08 57.0916.70 12.695.04 32.597.42

16 CBS112.17 13.596.14 30.7911.24 5.394.11 15.393.86

17 CBS207.252 0.090.00 3.793.22 0.090.00 3.592.38

18 CBS339.39 9.092.00 26.597.26 9.093.92 20.093.56

1Mean9standard deviation.2All the means of this isolate are significantly different with other isolate means within the same column atthe 5% level.



15A

Table 3. Differences in size and number of sclerotia of Sclerotinia minor isolates.

Length/Width2

MCG1 Isolate # PDA4 OAM5Sclerotia #/Plate3

PDA

1 IMI344141 1.0590.136 e7 1.1190.20 b 92.392.08 k8

LRC2104 1.1590.28 abcde 1.1490.23 b 461.3927.01 defg

ATCC44236 1.2590.30 abcd 1.1590.25 b 622.0963.91 abc

Sm44 1.1290.17 abcde 1.1590.27 b 697.0947.00 ab

Sm66 1.2590.29 abcd 1.2390.34 b 741.3945.54 a

2 S96-22 1.1790.23 abcde 1.2090.34 b 464.0914.00 defg

S96-138 1.1590.26 abcde 1.1490.23 b 376.0999.78 gh

S96-250 1.2590.28 ab 1.2290.25 b 535.3960.52 bcdef

S97-75 1.2590.30 abc 1.1690.28b 215.7914.36 j

3 R21 1.1190.21 de 1.2090.33 b 244.0921.28 ij

R23 1.1190.18 bcde 1.1290.40 b 344.3957.01 ghi

4 R22 1.0890.16 de 1.1790.34 b 301.7964.26 hij

R25 1.1490.27 cde 1.1290.28 b 433.7951.19 fg

5 BRIP28139 1.1290.17 abcde 1.1190.18b 448.094.36 efg

K4205 1.1290.21 bcde 1.2090.27 b 376.3934.43 gh

6 VPRI1671 1.290.25abcde 1.2290.28 b 620.0946.87 abc

VPRI1285 1.1290.23 de 1.2290.36b 352.3923.18 ghi

7 BPIC1681 n.a. n.a. 0.090.00 l

BPIC1949 1.3090.47 abcd 1.3090.37 b 612.0913.23 abcd

8 R24 1.1590.23 abcde 1.2290.31 b 224.3918.58 j

9 JW1 1.0790.20 e 1.1590.28 b 692.7928.11 ab

10 SMRF02 1.2890.31 ab 1.1590.27 b 353.7965.39 ghi

11 TH1C 1.15990.24 abcde 1.1690.25 b 519.3935.23 cdef

12 LU286 1.1790.25 abcde 1.4690.42 a 450.3953.69 efg

13 LK1 1.2190.28 abcde 1.2890.59 ab 589.3925.17 abcde

14 MUCL38484 1.2990.30 a 1.2190.29 b 576.3969.47 abcdef

15 S94-001 1.1290.19 abcde 1.1390.19 b 594.3931.66 abcde

16 CBS112.17 n.a. n.a. 0.090.00 l

17 CBS207.25 n.a. n.a. 0.090.00 l

18 CBS339.39 1.2890.35 abc 1.2390.29 b 45.097.94 k

1Mycelial compatibility groups.2The data were subjected to Kruskal�Wallis one-way ANOVA on ranks. Twenty-five individual sclerotia�3 replications through time for each isolate.

3The data were subjected to square root transformation to achieve normality then analyzed by one-wayANOVA. Average of three replications through time.

4Potato dextrose agar.5Oat meal agar.6Average9Standard deviation.7Within each column, means with common letters are not significantly different at P�0.05 according toTukey’s test.

8Isolate IMI344141 was significantly different from isolate CBS339.39 at P�0.006 according to the pairedt-test (SigmaStat for Windows version 2.03 1997).

9n.a., no sclerotia formed.



15A

S. sclerotiorum. Length/width ratio reflects sclerotia shape which was more

homogeneous among isolates on OMA than on PDA and ranged from 1.05 to 1.3

on PDA indicating a spherical to oblong shape.

A major distinguishing feature of isolate IMI 344141 is the low number ofsclerotia (Table 3). Sclerotia production of this isolate is significantly different from

all of the S. minor isolates tested including CBS339.39 (near S. sclerotiorum) which

produced significantly less number of sclerotia than IMI 344141 according to paired

t-test (Table 3).

Experiment 2. Mycelial compatibility groups

During the MCG test, incompatible pairings were recognized by the formation in aninteraction zone where mycelia from each isolate stopped growing. Accumulation of

the red food color in the hyphal tips formed a red line between incompatible pairings

at day 7 (Figure 1), whereas compatible isolates had fused mycelia associated with

abundant sclerotia in the fusion zone. There were 435 pairings between the 30

isolates with only 21 combinations showing a compatible reaction (4.8% of all the

combinations). Consistent pairing results were obtained in the replications of each

isolate. Mycelial compatible isolates were placed in the same MCG. Eighteen

mycelial compatibility groups (MCGs), seven multi-member (consisting of two tofive isolates each) and 11 single-member (R24, JW1, TH1C, SMRF02, LU286, LK1,

MUCL38484, S94-001, CBS112.17, CBS207.25, and CBS339.39) MCGs were

identified. Each MCG was assigned a number (MCG1-MCG18) (Table 1). MCG1

included the IMI 344141 isolate and four other isolates, MCG2 had four isolates

from Korea, MCG3 to MCG7 contained two isolates each, and MCGs 8-18 had one

isolate each. All self pairings were compatible. In all incompatible reactions, sclerotia

were not formed at the interaction zone (Figure 1). All compatible reactions

occurred within the first week of incubation. Because of the slow growth of isolateCBS207.25 (Table 2), pairing plates were incubated for 2 weeks and all pairings with

this isolate were incompatibile.

Experiment 3. Oxalic acid production

Oxalic acid was detected in the culture filtrate of all the studied S. minor isolates.

HPLC analysis revealed no significant differences in OA production among the

members of each of the MCGs (Figure 2), while there were some differences betweendifferent isolates of different MCGs. Maximum OA production was measured for

isolate S96-250 belonging to MCG 2, while minimum production was from isolates

R21 and CBS207.25.

Experiment 4. Isolate virulence

After 24 h, lesion diameters on detached dandelion leaves were variable among the

isolates and no lesions had been induced by nine isolates (Table 4). After 48 h, the S.minor isolates were classified into four categories: highly virulent (mean lesion

diameter ]0 mm), virulent (20�30 mm), moderately virulent (10�20 mm), and

hypovirulent (B10 mm). Isolate IMI 344141 fell within the moderately virulent

category with 26 mm lesion diameter, but the barley based formulation demonstrated



15A

high virulence without significant differences from the other highly virulent isolates

(Table 5). Although isolate TH1C ranked the fourth of the highly virulent isolates

using the mycelial disc method, it was the least virulent using the barley based

formulation.

Using dandelion plants, isolate IMI 344141 was one of the four highest isolates in

exerting significant above ground damage either 5 days or 3 weeks post application

(Figure 3). However, it was significantly ranked the first among all isolates in

reducing above and below soil biomass of dandelion (Figure 4). Moreover, the high

virulence of isolate IMI 344141 resulted in the least amount of re-growth and the

Figure 1. Pairings of Sclerotinia minor isolates demonstrating representative hyphal

interactions 7 days after inoculation (except C). (A) Isolate R24�BPIC1949, incompatible,

colony reverse. (B) Colony surface. (C) Isolate ATCC44236�CBS207.25, incompatible. (D)

Isolate Sm66�ATCC44236, compatible reaction. (E) Isolate Sm66�LRC2104, compatible

reaction. (F) Self-self reaction.



15A

lowest survival of dandelion among isolates after the first and the second

applications (Figure 5).

Descriptors distinguishing IMI 344141

IMI 344141 can be phenotypically distinguished from the other tested S. minor

isolates by vegetative compatibility testing and counting sclerotia produced on

standard 9-cm diameter PDA plates. IMI 344141 mycelia are white and fluffy and

sclerotia initials and sclerotia are formed on PDA agar in 96 and 144 h, respectively.

The sclerotia of IMI 344141 are spherical, black in color with a very rough surface.

Sclerotia are scattered on the surface of the plates rather than peripheral. IMI

344141 produces less than 100 sclerotia/9-cm diameter PDA plate.

Discussion

Variation within a worldwide collection of 30 isolates of S. minor was studied based

on culture morphology, mycelial compatibility and virulence. Two growth media,

PDA and OMA, were used to study the different culture characteristics of the 30

isolates including growth rates, colony type, mycelial color, time needed by each

isolate to form sclerotia, mode of sclerotia formation in addition to sclerotial

morphology counts and measurement and exudate formation. Differences in the

growth rate of all isolates were recorded on both media, where it was slower on OMA

than on PDA. Isolates showed different modes of sclerotia formation on both media,

where it was mostly concentric on OMA but peripheral or scattered on PDA.

abcd

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cde

e

bcde

de

abcd

abc

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abcd

e

abcd

eab

cde

de

dede

bcde

fab

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Sclerotinia minor isolates

Oxa

lic a

cid

conc

entra

tion

(mg

dl -1

)

0

25

50

75

100

125

150

175

200

225

250

275

S96-

138

S96-

22

SMR

F02

S94-

001

BPI

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49

S96-

250

LU28

6

MU

CL3

8484

S97-

75

VPR

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1

LK1

R24

JW1

TH1C

R21

R23

R22

R25

CB

S 11

2.17

CB

S207

.25

CB

S339

.39

BPI

C16

81

BR

IP28

139

VPR

I128

5

K42

05

IMI 3

4414

1

ATC

C44

236

Sm44

LRC

2104

Sm66

Figure 2. Oxalic acid production by different Sclerotinia minor isolates in potato dextrose

broth filtrate. Error bars refer to standard errors. Bars with common letters are not

significantly different at P�0.05 according to Tukey’s test.



15A

Whereas the use of the two growth media revealed these differences, it did not reveal

basic differences that we could rely on to differentiate between different isolates.

Smaller sclerotia sizes were measured for most isolates on OMA than on PDA.

Significant differences were found among isolates for sclerotia measurements on

both media where S94-001 and Sm66 had the smallest sclerotia on PDA and OMA,

respectively. CBS339.39 produced the largest sclerotia on both media. The culture

Table 4. Means and standard deviations (9 replicates) of the lesion diameters caused 24 and

48 h post inoculation of Sclerotinia minor isolates on detached dandelion leaves using 2-mm

mycelial discs.

Lesion diameter (mm)

MCG Isolate 24 h 48 h

1 IMI 344141 8.194.7 26.095.62

1 LRC2104 11.091.2 30.595.11

1 ATCC44236 4.295.1 25.996.92

1 Sm44 9.693.9 26.892.92

1 Sm66 11.49.9 28.997.72

2 S96-22 11.391.3 30.694.31

2 S96-138 12.791.7 37.196.91

2 S96-250 0.090.0 4.696.7

2 S97-75 10.792.4 30.194.71

3 R21 3.394.0 24.895.42

3 R23 12.391.0 35.393.41

4 R22 6.395.2 24.197.32

4 R25 7.294.4 18.392.43

5 K4205 11.191.2 28.695.52

5 BRIP28139 0.090.0 1.995.54

6 VPRI1671 0.090.0 13.195.63

6 VPRI1285 0.090.0 0.992.74

7 BPIC1681 0.090.0 0.090.04

7 BPIC1949 0.892.3 12.394.93

8 R24 0.090.0 0.090.04

9 JW1 4.095.0 18.098.83

10 SMRF02 12.390.7 35.094.11

11 TH1C 10.892.3 32.395.01

12 LU286 7.494.5 30.395.21

13 LK1 3.694.30 29.698.12

14 MUCL38484 9.695.7 28.597.72

15 S94-001 13.291.3 33.393.21

16 CBS 112.17 0.090.0 8.896.14

17 CBS207.25 0.090.0 0.090.04

18 CBS339.39 0.09 0.0 0.090.04

1Highly virulent ]30 mm.2Virulent (20�30 mm).3Moderately virulent (10�20 mm).4Hypo-and avirulent (B10 mm).



15A

Table 5. Means and standard deviations (15 replicates) of the lesion diameters caused 24 and

48 h post inoculation of the most aggressive Sclerotinia minor isolates on detached dandelion

leaves using colonized barley grits.

Lesion diameter (mm)

Isolate 24 h 48 h

IMI 344141 10.195.5 a 28.2911.4 a

R23 8.395.9 a 30.2910.0 a

LRC2104 4.894.7 ab 22.995.7 ab

SMRF02 8.494.7 a 27.5911.4 ab

TH1C 1.292.67 b 17.398.2 b

S94-001 6.295.77 ab 25.497.7 ab

S96-22 6.595.3 ab 21.9910.7 ab

S96-138 10.192.7 a 28.395.5 a

S97-75 6.794. 7 ab 28.194.7 a

LU286 8.494.7 a 27.5911.4 ab

Within each column, means with similar letters are not significantly different at P�0.05 according toTukey’s test.

Sclerotina minor isolatesIM

I3441

41

S96-18

3

S97-75

S94-00

1

S96-22

SMRF02TH1C LR

CLU

286

R23

Abo

ve-g

roun

d da

mag

e (%

)

0

20

40

60

80

100

120 5 days post application 3 weeks post application

aab

abcab

abc

a

c

ab

bcabc

m

n

o

mn mnmn

non nn

Figure 3. Above ground damage to dandelion caused by different Sclerotinia minor isolates.

Error bars represent the standard errors of the means (average of 16 plant replicates). Within

each post application time, bars with a common letter are not significantly different at P�0.05

according to Tukey’s test.



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characteristics and large dimensions of the sclerotia of this isolate suggest that

CBS339.39 was misclassified and is likely closer to S. sclerotiorum. Spherical to

oblong sclerotial shape was a reflection of length/width ratio measurements, which

was more homogeneous on OMA than on PDA. Woodard and Simpson (1993)

reported a range of 1000�3000 sclerotia per plate on PDA for S. minor isolates from

peanut. Our range of sclerotia production was lower than this even for those isolates

isolated from peanut plants (R21, R23, SMRF02). Most isolates produced 200�700sclerotia per PDA plate, whereas two isolates: IMI 344141 and CBS339.39 produced

3.0

2.5

2.0

1.5

1.0

0.5

0.0

Bio

mas

s (g

pla

nt -1

)

0.0

0.5

1.0

1.5

2.0

2.5

3.0

IMI3

4414

1

S96

-183

S97

-75

S94

-001

S96

-22

SM

RF0

2

TH1C

LU28

6

R23

LRC

cont

rol

30

2.5

2.0

1.5

1.0

0.5

Above-groundRoots

a

b

c

bb

bb

b

b

bb

a

b

bbc

cd

bbcbc bc bc

Figure 4. Effects of different Sclerotinia minor isolates on the above- and below-ground

biomass of dandelion after two consecutive spot applications of 0.2 g/plant of S. minor

colonized barley granules. Error bars represent the standard errors of the means (average of 16

plant replicates). Within each group, bars with a common letter are not significantly different

at P�0.05 according to Tukey’s test.



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significantly fewer sclerotia per plate on PDA than all sclerotia-producing isolates.

Non-sclerotia producing isolates were also part of this study.

Diversity in fungal populations can be measured by MCGs (Leslie 1993).

Mycelial compatibility in different Sclerotinia species, other than S. minor, has been

comprehensively studied. The present study demonstrates for the first time the

occurrence of MCG diversity within S. minor. Results from this study showed that,

this worldwide collection of S. minor isolates is represented by 18 MCGs, seven of

them as multi-member groups. Of the tested isolates, 36.6% were unique isolates that

could not be assigned to any of the eight multi-member MCGs indicating numerous

single-member MCGs may exist. MCG Diversity is the ratio between the number of

MCGs and the total number of isolates (Viji et al. 2005). Our results demonstrate

high diversity (0.6, highest diversity is 1) among the tested isolates of S. minor which

agree with the findings for the closely related species S. sclerotiorum (Kohn et al.

1991). This high diversity among MCGs could be correlated with the wide host range

reported for S. minor (Melzer et al. 1997; Hollowell et al. 2003).

The recovery of an MCG of S. minor in diverse geographical areas (MCG1) or

hosts (MCG4, MCG5 and MCG6) could be attributed to spread by agricultural

practices and other human activities through contaminated soil, crop material, and

equipment (Harlton, Levesque, and Punja 1995; Punja and Sun 2001; Cilliers,

Pretorius, andVanWyk 2002; Durman et al. 2003; Atallah, Larget, Chen, and Johnson

2004). The occurrence of certainMCGs likeMCG3andMCG7 in a specific geographic

region oron the same host species has been reported forSclerotium rolfsii (Nalim, Starr,

Woodard, Segner, and Keller 1995; Punja and Sun 2001; Cilliers et al. 2002).

Virulence assays could help in subdividing populations of fungi into different

groups and isolates within a MCG could have similar virulence (Kull and Pedersen

2004; Viji et al. 2005). In our study, however, results from the virulence tests did not

show a distinct pattern of relationship between virulence and MCG except for three

Sclerotinia minor isolatesIM

I3441

41

S96-18

3

S97-75

S94-00

1

S96-22

SMRF02TH1C LR

CLU

286

R23

Dan

delio

n su

rviv

al (%

)

0

20

40

60

80

100

IMI34

4141

S96-18

3

S97-75

S94-00

1

S96-22

SMRF02TH1C LR

CLU

286

R230

20

40

60

80

100

Partial damageRe-growth after 100% above-ground damage

b

a aa

a

a

a aa

a a

aa

a

a a

b

bc

c

a

A B

Figure 5. Effects of different Sclerotinia minor isolates on re-growth from roots and

dandelion survival. (A) Three weeks after the first application of 0.2 g plant�1 of S. minor

barley based formulation; (B) 3 weeks after the second application of the same rate. Mean

values represent averages of 16 plant replicates. Within each graph, bars with a common letter

are not significantly different at P�0.05 according to Tukey’s test.



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isolates, S96-22, S96-138 and S97-75, from MCG2 and the isolates of MCG1 where

the lesion diameter ranged between 26 and 30.5 mm. Positive correlation between

virulence in vitro and mycelial growth was not always applicable, since many of the

studied isolates with high growth rate on PDA (Table 2) were hypovirulent and thiscontradicts earlier reports where positive correlation was observed between mycelial

growth and aggressiveness in vitro for isolates of S. sclerotiorum (Boland and Hall

1992; Durman et al. 2003).

In our study, virulence tests on detached dandelion leaves were performed using

two types of inocula, mycelial discs and the barley based formulation. In the mycelial

discs study, measurements were taken 24 and 48 h after inoculation and relative

virulence of the different isolates was ranked according to lesion size after 48 h. IMI

344141 has strong bioherbicidal activity against dandelion (Abu-Dieyeh et al. 2005;Abu-Dieyeh and Watson 2006) and appeared in the moderate virulent group of

isolates. However, when the barley-based formulations of S. minor were tested

on detached dandelion leaves, IMI 344141 was one of the most virulent isolates

(Table 5). When the barley based formulations of the 10 most virulent isolates were

applied onto dandelion plants, IMI 344141 caused the greatest biocontrol effect in

reducing above and below ground biomass (Figures 3�5). These results suggest

caution in relying on detached leaves in virulence assessment of S. minor.

Earlier studies attempted to positively correlate virulence with levels of oxalicacid produced by the fungi Cryphonectria parasitica (Vinnini et al. 1993), Sclerotium

rolfsii (Bateman and Beer 1965; Kritzman et al. 1977), and Sclerotinia sclerotiorum

(Marciano, Magro, and Favaron 1989; Godoy et al. 1990; Boland and Hall 1992;

Zhou and Boland 1999). Conflicting data regarding differences in oxalic acid

production by hypovirulent and virulent fungal isolates have been reported (Havir

and Anagnostakis 1983; Bennett and Hindal 1989). In our study, there were no

significant differences in production of OA among isolates of the same MCG and

there was no correlation between OA production and virulence or growth rate onsolid media since many of the high OA producing isolates had slow growth and were

hypovirulent.

IMI 344141 was clearly distinguished from all other S. minor isolates tested based

on the number of sclerotia produced on PDA. IMI 344141 produced fewer than 100

sclerotiaper platewhile otherS.minor isolates producedmore than 200 andupwards of

700 sclerotia per plate. In conclusion, groups based on morphological or pathological

characters could not be easily related to mycelial compatibility groups and without

molecular marker genotyping, MCGs alone do not suffice to completely characterizepopulations. To this end, molecular characterization of IMI 344141 is ongoing.

Acknowledgements

The authors are grateful to Miron Teshler for his help in various aspects of the project.Financial support from the Natural Sciences and Engineering Research Council of Canada(NSERC) Idea to Innovation (I2I) grant and Sarritor Inc. are gratefully acknowledged. Thefollowing graciously loaned S. minor isolates: G. Abawi, Cornell University, Geneva, USA; G.Boland, University of Guelph; J. Whipps, Warwick Horticulture Research International, UK;H. Huang, Agriculture and Agri-food Canada, Brandon; P. Phipps, Virginia Tech; S. O’Neill,Queensland Dept. Plant Industry, Brisbane; Tamrika Hind-Lanoiselet, Wagga WaggaAgricultural Institute, NSW; S. Morley, Herbarium VPRI, Victorian Department of PrimaryIndustries, Knoxfield, Victoria, Australia; K. Elena, Benaki Phytopathological Institute,Athens, Greece; Alison Stewart, Lincoln University, Christchurch, New Zealand; Levente



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Kiss, Hungarian Plant Protection Institute, Budapest; P. Charue, Belgian CoordinatedCollections of Microorganisms, Louvain; J. Tatnell and M. Fuhlbohm, Australian Depart-ment of Plant Industry, Kingaroy, QSLD; W.G. Kim, National Institute of AgriculturalScience and Technology, Suwon, Republic of Korea; F. Snippe-Claus, Centraalbureau voorSchimmelcultures, Utrecht, The Netherlands.

References

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Abu-Dieyeh, M.H., and Watson, A.K. (2006), ‘Effect of Turfgrass Mowing Height onBiocontrol of Dandelion with Sclerotinia minor’, Biocontrol Science and Technology, 16,509�524.

Abu-Dieyeh, M.H., and Watson, A.K. (2007a), ‘Grass Over-seeding and a Fungus Combineto Control Taraxacum officinale’, Journal of Applied Ecology, 44, 115�124.

Abu-Dieyeh, M.H., and Watson, A.K. (2007b), ‘Efficacy of Sclerotinia minor for DandelionControl: Effect of Dandelion Accession, Age and Grass Competition’, Weed Research, 4,63�72.

Abu-Dieyeh, M.H., and Watson, A.K. (2007c), ‘Population Dynamics of Broadleaf Weeds inTurfgrass as Influenced by Chemical and Biological Control Methods’, Weed Science, 55,371�380.

Abu-Dieyeh, M.H., Bernier, J., and Watson, A.K. (2005), ‘Sclerotinia minor Advances Fruitingand Reduces Germination in Dandelion (Taraxacum officinale)’, Biocontrol Science andTechnology, 15, 815�825.

Atallah, Z.K., Larget, B., Chen, X., and Johnson, D.A. (2004), ‘High Genetic Diversity,Phenotypic Uniformity and Evidence of Outcrossing in Sclerotinia sclerotiorum in theColumbia Basin of Washington State’, Phytopathology, 94, 737�742.

Bateman, D.F., and Beer, S.V. (1965), ‘Simultaneous Production and Synergistic Action ofOxalic Acid and Polygalactronase during Pathogenesis by Sclerotium rolfsii’, Phytopathol-ogy, 55, 204�211.

Bennett, A.R., and Hindal, D.F. (1989), ‘Mycelial Growth and Oxalate Production byFive Strains of Cryphonectria parasitica in Selected Liquid Culture Media’, Mycologia, 81,554�560.

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Briere, S.C., Watson, A.K., and Paulitz, T.C. (1992), ‘Evaluation of Granular Sodium AlginateFormulations of Sclerotinia minor Jagger, as Potential Biocontrol Agents of Turfgrass WeedSpecies’, Phytopathology, 82, 1081.

Briere, S.C., Watson, A.K., and Hallett, S.G. (2000), ‘Oxalic Acid Production and MycelialBiomass Yield of Sclerotinia Minor for the Formulation Enhancement of a Granular TurfBioherbicide’, Biocontrol Science and Technology, 10, 281�289.

Callahan, F.E., and Rowe, D.E. (1991), ‘Use of Host-Pathogen Interaction System to TestWhether Oxalic Acid is the Sole Pathogenic Determinant in the Exudates of Sclerotiniatrifoliorum’, Phytopathology, 81, 1546�1550.

Cilliers, A.J., Pretorius, Z.A., and Van Wyk, P.S. (2002), ‘Mycelial Compatibility Groups ofSclerotium rolfsii in South Africa’, South Africa Journal of Botany, 68, 389�392.

Ciotola, M., Wymore, L., and Watson, F (1991), ‘Sclerotinia, a Potential Mycoherbicide forLawns’, WSSA Abstracts 30, Weed Science Society of America Annual meeting, Feb. 4�7,1991, Louisville, KY, USA.

Durman, S.B., Menendez, A.B., and Godeas, A.M. (2003), ‘Mycelial Compatibility Groups inBuenos Aires Field Population of Sclerotinia sclerotiorum (Sclerotiniaceae)’, AustralianJournal of Botany, 51, 421�427.

Godoy, G., Steadman, J.R., Dickman, M.B., and Daur, R. (1990), ‘Use of Mutants toDemonstrate the Role of Oxalic Acid in Pathogenicity of Sclerotinia sclerotiorum onPhaseolus vulgaris’, Physiological and Molecular Plant Pathology, 37, 179�191.

Harlton, C.E., Levesque, C.A., and Punja, Z.K. (1995), ‘Genetic Diversity in Sclerotium(Athelia) rolfsii and Related Species’, Phytopathology, 85, 1269�1281.



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Havir, E., and Anagnostakis, S.L. (1983), ‘Oxalate Production by Virulent but notHypovirulent Strains of Endothia parasitica’, Physiological Plant Pathology, 23, 369�376.

Hollowell, J.E., and Shew, B.B. (2003), ‘Evaluating Isolate Aggressiveness and Host Resistancefrom Peanut Leaflet Inoculations with Sclerotinia minor’, Plant Disease, 87, 402�406.

Hollowell, J.E., Smith, M.R., and Shew, B.B. (2001), ‘Oxalic Acid Production by Nine Isolatesof Sclerotinia minor’, Proceedings of American Peanut Research and Education Society, 33, 24.

Hollowell, J.E., Shew, B.B., Cubeta, M.A., and Wilcut, J.W. (2003), ‘Weed Species as Hosts ofSclerotinia minor in Peanut Fields’, Plant Disease, 87, 197�199.

Kohn, L.M., Carbone, I., and Anderson, J.B. (1990), ‘Mycelial Interactions in Sclerotiniasclerotiorum’, Experimental Mycology, 14, 255�267.

Kohn, L.M., Stasovski, E., Carbone, I., Royer, J., and Anderson, J.B. (1991), ‘MycelialIncompatibility and Molecular Markers Identify Genetic Variability in Field Populations ofSclerotinia sclerotiorum’, Phytopathology, 81, 480�485.

Kritzman, G., Chet, I., and Henis, Y. (1977), ‘The Role of Oxalic Acid in the PathogenicBehavior of Sclerotium rolfsii Sacc.’, Experimental Mycology, 1, 280�285.

Kull, L.S., and Pedersen, W.L. (2004), ‘Mycelial Compatibility and Aggressiveness ofSclerotinia sclerotiorum’, Plant Disease, 88, 325�332.

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Leslie, J.F. (1996), ‘Fungal Vegetative Compatibility Promises and Prospects’, Phytoparasitica,24, 3�6.

Lumsden, R.D. (1979), ‘Histology and Physiology of Pathogenesis in Plant Diseases Causedby Sclerotinia spp.’, Phytopathology, 69, 890�896.

Marciano, P., Magro, P., and Favaron, F. (1989), ‘Sclerotinia sclerotiorum Growth and OxalicAcid Production on Selected Culture Media’, FEMS Microbiology Letters, 61, 57�60.

Melzer, M., Smith, E., and Boland, G. (1997), ‘Index of Plant Hosts of Sclerotinia minor’,Canadian Journal of Plant Pathology, 19, 272�280.

Nalim, F.A., Starr, J.L., Woodard, K.E., Segner, S., and Keller, N.P. (1995), ‘MycelialCompatibility Groups in Texas Peanut Field Populations of Sclerotium rolfsii’, Phyto-pathology, 85, 1507�1512.

Noonan, M.P., Glare, T.R., Harvey, I.C., and Sands, D.C. (1996), ‘Genetic Comparison ofSclerotinia Isolates from New Zealand and USA’, Proceedings 49th New Zealand PlantProtection Conference, New Zealand Plant Protection Society (Inc.). pp. 126�131.

Pest Management Regulatory Agency (PMRA), (2001), ‘Guidelines for the Registration ofMicrobial Pest Control Agents and Products’ Regulatory Directive, DIR2001-02, PestManagement Regulatory Agency, Health Canada, Ottawa, ON, 99 pp.

Pest Management Regulatory Agency (PMRA), (2007), ‘Evaluation Report Sclerotinia minorstrain IMI 344141’, Available from: Publications Internet: [email protected],Pest Management Regulatory Agency, Health Canada, Ottawa, ON, 50 pp.

Punja, Z.K., and Grogan, R.G. (1983), ‘Hyphal Interaction and Antagonism among FieldIsolates and Single � Basidiospore Strains of Athelia (Sclerotium rolfsii)’, Phytopathology,73, 1279�1284.

Punja, Z.K., and Sun, L. (2001), ‘Genetic Diversity among Mycelial Compatibility Groups ofSclerotium rolfsii (teleomorph Athelia rolfsii) and S. delphinii’, Mycological Research, 105,537�546.

Riddle, G., Burpee, L., and Boland, G. (1991), ‘Virulence of Sclerotinia sclerotiorum andS. minor on Dandelion (Taraxacum officinale)’, Weed Science, 39, 109�118.

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Schnick, P., Stewart-Wade, S., and Boland, G. (2002), ‘2,4-D and Sclerotinia minor to ControlCommon Dandelion’, Weed Science, 50, 173�178.

Stewart-Wade, S.M., Green, S., Boland, G.J., Teshler, M.P., Teshler, I.B., Watson, A.K.,Sampson, M.G., Patterson, K., DiTommaso, A., and Dupont, S. (2002), ‘82. Taraxacumofficinale Weber, Dandelion (Asteraceae)’, in Biological Control Programs in Canada, 1981�2000, eds. P.G. Mason and J.T. Huber, New York: CABI Publishing.



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Viji, G., Uddin, W., O’Neill, N.R., Mischke, S., and Saunders, J.A. (2004), ‘Genetic Diversityof Sclerotinia homoeocarpa Isolates from Turf Grasses from Various Regions in NorthAmerica’, Plant Disease, 88, 1269�1276.

Willetts, H., and Wong, J. (1980), ‘The Biology of Sclerotinia sclerotiorum, S. trifoliorum andS. minor with Emphasis on Specific Nomenclature’, The Botanical Review, 46, 101�165.

Woodard, K.E., and Simpson, C.E. (1993), ‘Characterization of Growth and SclerotialProduction of Sclerotinia minor Isolated from Peanut in Texas’, Plant Disease, 77, 576�579.

Zhou, T., and Boland, G.J. (1999), ‘Mycelial Growth and Production of Oxalic Acid byVirulent and Hypovirulent Isolates of Sclerotinia sclerotiorum’, Canadian Journal of PlantPathology, 21, 93�99.



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10. SCLEROTINIA MINOR–BIOCONTROL TARGET OR AGENT?

ALAN WATSON* Department of Plant Science, McGill University, Sainte-Anne-de-Bellevue, Quebec, Canada

Abstract. Sclerotinia minor is the causal agent of several important crop diseases including lettuce drop and Sclerotinia blight of peanut. Extensive search for biocontrol of the Sclerotinia diseases has culminated in the commercialization of Coniothyrium minitans. Sclerotinia minor is also an effective bioherbicide that can be effectively and safely used to control broadleaf weeds in turf environments.

Keywords: host range, crop pathogen, virulence, dissemination.

10.1 Sclerotinia minor- The Crop Pathogen, a Target for Biological Control

Sclerotinia minor Jaggar is a soil borne Discomycetes fungus characterized by small (0.5-2.0 mm), irregular sclerotia that germinate by eruptive growth of mycelium and colonize susceptible plant tissues.1,2 S. minor is closely related to S. sclerotiorum (Lib.) de Bary and S. trifoliorum Erikss. These species are serologically related with S. sclerotiorum a tetraploid form of S. minor whilst S. trifoliorum a hybrid with part of the genome contributed by S. minor.3 In contrast to S. sclerotiorum, apothecia and ascospore production in S. minor is very rare in the field and has not been recorded to occur in North America4 and only one report from New Zealand5 of their natural occurrence. Various workers1,6 have concluded that ascospores are unimportant in the epidemiology of S. minor caused disease. Sclerotinia minor has been known to occur in North America prior to 1900 and has been the target of extensive research with results widely published1,2,4,6 in the scientific literature.

* To whom correspondence should be addressed. e-mail: [email protected]

1 Novel Biotechnologies for Biocontrol Agent Enhancement and Management © 2007 Springer. Printed in the Netherlands.

______

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SCLEROTINIA BIOHERBICIDE 2

1.1. DISTRIBUTION AND HOST RANGE

S. minor is distributed worldwide, except for the warm tropics, and occurs on many plant species.7 Most susceptible species are dicotyledonous with only three monocotyledonous plants, asparagus, tulip, and banana, reported as hosts. S. minor is not a serious pathogen on most plants as economic losses to S. minor in the United States have only occurred in lettuce8 and peanuts9 and in eastern Canada,10 only on lettuce. The most economically important diseases caused by S. minor are lettuce drop, Sclerotinia blight of peanut, and Sclerotinia stem rot of sunflower. S. minor is not pathogenic on any species of the grass family Poaceae.

1.2. LIFE CYCLE

Dispersal and transmission of the disease is exclusively by the direct contact with germinating sclerotia to produce infective hyphae which colonize plants and eventually produce more sclerotia which are returned to the soil.4,11 Sclerotia must be within 2 cm of the taproot of lettuce and 8 cm of the soil surface to cause disease.12 Plant-to-plant spread between diseased and healthy plants can occur by direct contact with infected tissue.4 Infection of lettuce with S. minor can occur at the soil line through lower senescent leaves or below ground to a depth of 10 cm through root tissues.8 Favourable conditions for germination and infection include temperatures of 17-21C and relative humidity greater than 95 percent. Infection occurs by mycelium arising from sclerotia or infected plant debris. Plants may be attacked at any stage from seedling to maturity. Under moist and cool conditions the fungus rapidly invades the tissues of the host in which a light brown, watery rot develops and a white, cottony-like mycelium grows over the infected tissues. Stunting, premature ripening and sudden collapse of the host are common symptoms of infection. After several days of mycelium growth, small, compact bodies develop either on the surface of the host or in cavities within it. These aggregates of mycelium are young resting vegetative structures (sclerotia). At first they are white, but when mature are black in color. Large numbers of sclerotia accumulate in plant debris and in soil where they can remain dormant for long periods. Alternatively they may germinate after a short resting period.

The S. minor inoculum of lettuce drop disease is sedentary and spread between commercial fields is slow and restricted.8,13 After harvest, lettuce residues infested by S. minor, including those with sclerotia already formed are disked into the soil where the inoculum remains dormant until the next planting. Disease is often in clumped or aggregated distribution patterns within infested fields. Several modes of dissemination of S. minor have

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been proposed; mycelia infected seed, seed contaminated with sclerotia, movement of infested soil by machinery, winter annual weed species acting as reservoirs in rotational systems and passage through animals fed or bedded with peanut plant material.14 Viable sclerotia of S. minor were recovered from fecal and ruminal samples of heifers fed infested peanut hay.15

1.3. SURVIVAL AND PERSISTENCE

Soil moisture and temperature, sclerotial position and duration in the soil, sclerotia shape, soil gases and chemicals, microbial activity, nutrition, and other factors are known to affect survival and germination of sclerotia.6,11

There have been reports16 of the sclerotia of Sclerotinia species surviving in the soil for 4-5 years while others report17 rapid decline in sclerotia survival with few surviving into the following year. In soil box and field trials in New Zealand, only 22% of the S. minor sclerotia could be recovered after 3 months, and only 2% (which were 50 % viable) were recovered after 11 months.17 In water saturated soil, sclerotia of S. minor disintegrate or fail to germinate within 8 weeks.18

1.4. NATURAL CONTROL OF SCLEROTINIA MINOR

The biological component of the soil is the most important component affecting survival of S. minor sclerotia. Forty-six fungi, two bacteria, two insects, a mite, and a snail are reported as antagonists, mycoparasites or predators of Sclerotinia spp.11,16,19.. These organisms are thought to be responsible for the occurrence of “suppressive soils”. Several; including Coniothyrium minitans, Trichoderma harzianum, Teratosperma oligocladum, Talaromyces flavus, and Sporidesmium sclerotivorum have been evaluated as biocontrol agents to deal with Sclerotinia spp. in lettuce with varying degrees of success.4,18,20,21 CONTANS WG, a water dispersible granule formulation of Coniothyrium minitans is registered for the reduction/control of Sclerotinia sclerotiorum and Sclerotinia minor in agricultural soils in Europe and the United States (Chapter 12).

2. Sclerotinia spp. as bioherbicides

The severe and rapid necrosis caused by Sclerotinia sclerotiorum on a wide spectrum of broadleaf weeds has created interest in utilizing S. sclerotiorum as a biological agent to control weeds. S. sclerotiorum was field tested as a bioherbicide against Centaurea maculosa (spotted knapweed) in British Columbia in 1972 (Watson, unpublished). In Montana state wide trials in

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1982, S. sclerotiorum controlled 20 to 80% of Cirsium arvense (Canada thistle).22 This work was followed by the selection of non-sclerotia mutants23 incapable of producing ascospores, but virulence is linked to sclerotia formation. Amino acid auxotroph mutants24 were also developed in attempts to mitigate reproduction and persistence of a S. sclerotiorum bioherbicide. The pathogenicity of amended auxotrophic strains and wild strains of S. sclerotiorum were compared in a permanent pasture in New Zealand and the auxotrophic strains were less field fit than the wild strains.25

An elaborate risk analysis of using S. sclerotiorum for biological control of Cirsium arvense simulated dispersal of ascospores in a pasture.26 A ‘safety zone’ was determined to be the distance from a pasture undergoing biological weed control using S. sclerotiorum at which the concentration of dispersing ascospores has declined to that occurring naturally in the air. Regional variation in the width of ‘safety zones’ for sheep and dairy pasture treated with a S. sclerotiorum mycoherbicide have been quantified using climatic data and wind direction.27.

Interest in the weed control potential of S. minor was first reported in 199128,29 when several Sclerotinia species were compared. In one study28, a S. minor isolate was more virulent on dandelion than the S. sclerotiorum and S. trifoliorum isolates. Subsequently, this S. minor isolate (IMI 344141) became the focus of bioherbicide research. 30-34

3. Sclerotinia minor IMI 344141 “sensu stricto”- the bioherbicide

S. minor IMI 344141 was obtained from a lettuce field in Sherrington, Québec in 1983. The life cycle, mode of action, moisture and temperature requirements, and host range of S. minor IMI 344141 are not different from S. minor “sensu lato”. However, persistence, survival and dissemination are much different when S. minor IMI 344141 is employed as an integrated biocontrol product.

3.1. THE BIOHERBICIDE PRODUCT

Sclerotinia minor IMI 344141 is the active ingredient of SARRITOR, a bioherbicide proceeding towards registration as a Microbial Pest Control Product (MPCP) in Canada for the control of Taraxacum officinale (dandelion) and other broadleaf weeds in turfgrass. The fungus is cultured on ground barley and the bioherbicide granules are broadcast applied to weed infested turf. Favourable conditions for germination and infection include 15-24C temperatures and 95+ percent relative humidity. Disease develops quickly and complete kill of dandelion and other broadleaf weeds

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can be achieved within 7 days, about twice as fast as the standard chemical herbicide KillexTM. The product is compatible with normal lawn maintenance operations such as mowing, fertilization and irrigation.

3.2. SURVIVAL AND PERSISTENCE OF S. MINOR IMI 344141

Questions concerning persistence of the SARRITOR product and effect on turfgrass have been addressed in greenhouse and fields studies 30,32,33,. When applied to turfgrass, S. minor IMI 344141 rarely produces sclerotia (melanized survival structures) and these sclerotia do not survive over winter. Sclerotia formation is mainly associated with clumps of inoculum rather than infected weed tissue. Eruptive mycelial growth of S. minor IMI 344141 from the inoculum does not persist in the absence of a host and quickly decays within 10 days. Lettuce bioassays of treated field trials have revealed no infectivity of S. minor IMI 344141 in the turf environment four months after treatment. Field and greenhouse studies confirmed that turfgrass species are not susceptible to S. minor IMI 344141.32,33 Independent human health and environmental toxicology studies established that S. minor IMI 344141 is neither toxic nor pathogenic to non-target organisms. These data support MPCA registration and have been incorporated within the product submission to the Pest Management Regulatory Agency in Ottawa, Canada.

3.3. OFF TARGET MOVEMENT OF S. MINOR IMI 344141.

Sclerotinia minor IMI 344141 does not move off target. When applied, the granules settle down within the turf on or near the soil surface. Granules are not easily dislodged or dispersed from the point of application. SARRITOR granules have been applied to over 250 field research plots in Eastern Canada and there has been no occurrence of off-target movement expressed as disease on plants beyond plot borders.

3.4. WEED CONTROL EFFICACY OF SARRITOR

The Sclerotinia minor IMI 344141 bioherbicide provides effective control of dandelion and many other broadleaf weeds including broadleaf plantain (Plantago major), white clover (Trifolium repens), and field bindweed (Convolvulus arvensis). Under high weed infestation levels in the field, S. minor caused a greater initial reduction of dandelion density than did the herbicide during the two-weeks-post application period, although reductions were greater in herbicide treated plots by six weeks after application.32 Over the growing season, S. minor and the herbicide had similar

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suppressive effects on dandelion density except under low mowing height (3-5 cm). Sclerotinia minor IMI 344141 has no residual activity, thus a vigourous competitive grass sward enhances the efficacy of S. minor by minimizing dandelion seedling recruitment in vegetation gaps created by the complete removal of the dandelions.33,34 All life stages of dandelion from seeds to flowering plants are susceptible to the Sclerotinia minor IMI 344141 disease. Disease symptoms were identical on 14 different accessions of dandelion from Europe and North America and the above-ground and below-ground biomass were reduced by 94% and 96%, respectively with no difference among accessions. 34

Unlike most host specific, questionable virulence bioherbicide candidates that are being investigated worldwide, Sclerotinia minor IMI 344141 is a highly virulent, and broad spectrum. In addition to being an important crop pathogen, Sclerotinia minor can also provide effective broadleaf weed control in turfgrass.

References

1. H. R. Dillard and R. G. Grogan, Relationship between sclerotial spatial pattern and density of Sclerotinia minor and the incidence of lettuce drop. Phytopathology 75, 90-94 (1985).

2. H. J. Willets and J. A-J. Wong, The biology of Sclerotinia sclerotiorum, S. trifoliorum, and S. minor with emphasis on specific nomenclature. Bot. Rev. 46, 101-165 (1980).

3. S. W. Scott. Serological relationships of three Sclerotinia species. Trans. Brit. Mycol. Soc. 77, 674-676 (1981).

4. K. V. Subbarao, Progress toward integrated management of lettuce drop. Plant Dis. 82, 1068-1078 (1998).

5. B. T. Hawthorne, Observations on the development of apothecia of Sclerotinia minor Jagg. in the field. N. Z. J. Agr. Res. 19, 383-386 (1976).

6. J. J. Hao, K. V. Subbarao, and J. M. Duniway, Germination of Sclerotinia minor and S. sclerotiorum sclerotia under various soil moisture and temperature combinations. Phytopathology 93, 443-450 (2003).

7. M. S. Melzer, E. A. Smith and G. J. Boland, Index of hosts of Sclerotinia minor. Can. J. Plant Pathol. 19, 272-280 (1997).

8. S. Abawi and R. G. Grogan, Epidemiology of diseases caused by Sclerotinia species. Phytopathology 69, 899-904 (1979).

9. D. M. Porter and M. K. Buete, Sclerotinia blight of peanuts. Phytopathology 64, 263-264 (1974).

10. W. R. Jarvis, Sclerotinia minor as the cause of lettuce drop in southwestern Ontario. Can. Plant Dis. Sur. 65, 1 (1985).

11. P. B. Adams, Effects of soil temperature, moisture and depth on survival and activity of Sclerotinia minor, Sclerotium cepivorum and Sporidesmium sclerotivorum. Plant Dis. 71, 170-174 (1987).

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12. J. J. Hao and K. V. Subbarao, Comparative analyses of lettuce drop epidemics caused by Sclerotinia minor and S. sclerotiorum. Plant Dis. 89, 717-725 (2005).

13. E. D. Imolehin, R. G. Grogan and J. M. Duniway, Effect of temperature and moisture tension on growth, sclerotial production, germination, and infection by Sclerotinia minor. Phytopathology 70, 1153-1157 (1980).

14. J. E. Hollowell, G. G. Shaw, M. A. Cubeta and J. W. Wilcut, Weed species as hosts of Sclerotinia minor in peanut fields. Plant Dis. 87, 127-199 (2003).

15. H. A. Melouk, L. L. Singleton, F. N. Owens and C. N. Akem, Viability of sclerotia of Sclerotinia minor after passage through the digestive tract of a crossbred heifer. Plant Dis. 73:68-69 (1989).

16. B. Adams and W. A. Ayers, Ecology of Sclerotinia species. Phytopathology 69, 896-899 (1979).

17. B. J. R. Alexander and A. Stewart, Survival of sclerotia of Sclerotinia and Sclerotium spp in New Zealand horticultural soil. Soil Biol. Biochem. 26, 1323-1329 (1994).

18. E. D. Imolehin and R. G. Grogan, Factors affecting survival of sclerotia and effects of inoculum density, relative position, and distance of sclerotia from the host on infection of lettuce by Sclerotinia minor. Phytopathology 70, 1162-1167 (1980).

19. J. B. Coley-Smith and R. C. Cooke, Survival and germination of fungal sclerotia. Ann. Rev. Phytopath. 9, 65-92 (1971).

20. E. E. Jones and A. Stewart, Biological control of Sclerotinia minor in lettuce using Trichoderma species. Proc. 50th N. Z. Plant Prot. Conf. (1997) pp. 154-158.

21. H. J. Ridgway, N. Rabeendran, K. Eade and A. Steart, Application timing of Coniothyriun minitans A69 influences biocontrol of Sclerotinia minor in lettuce. N. Z. Plant Prot. 54, 89-92 (2001).

22. B. S. Brosten and D. C. Sands, Field trials of Sclerotinia sclerotiorum to control Canada thistle (Cirsium arvense). Weed Science 34, 377-380 (1986).

23. C. Miller, E. F. Ford and D. Sands, A nonnsclerotial pathogenic mutant of Sclerotinia sclerotiorum. Canadian Journal of Microbiology 35, 517-520 (1989).

24. R. V Miller, E. J. Ford, N. J. Zidack, and D. C. Sands, A pyrimidine auxotroph of Sclerotinia sclerotiorum for use in biological weed control. J. Gen. Microbiol. 135, 2085-2091 (1989).

25. I. C. Harvey, G. W. Bourdot, D. J. Saville, and D. C. Sands, A comparison of auxotrophic and wild strains of Sclerotinia sclerotiorum used as a mycoherbicide against Californian thistle (Cirsium arvense). Biocontrol Sci. Tech. 8, 73-81 (1998).

26. M. D. de Jong, G. W. Bourdot, G. A. Hurrell, D. J. Saville, H. J. Erbrink and J. C. Sadoks, Risk analysis for biological weed control – simulating dispersal of Sclerotinia sclerotiorum (Lib.) de Bary ascospores from a pasture after biological control of Cirsium arvense (L.) Scop. Aerobiologica 18, 211-111 (2002).

27. G. W. Bourdôt, D. Baird, G. A. Hurrell and M. D. De Jong. Safety zones for a Sclerotinia sclerotiorum-based mycoherbicide: Accounting for regional and yearly variation in climate. Biocontrol Sci. Tech.. 16, 345-358 (2006).

28. M. Ciotola, L. A. Wymore and A. K. Watson, Sclerotinia, a potential mycoherbicide for lawns. Weed Abst. 31, 81 (1991).

29. G. E. Riddle, L. L. Burpee and G. J. Boland, Virulence of Sclerotinia sclerotiorum and S. minor on dandelion. Weed Sci. 39, 109-118 (1991).

30. S. M. Stewart-Wade, S. Green, G. J. Boland, M. P. Teshler, I. B. Teshler, A. K. Watson, M. G. Sampson, K. Patterson, A. DiTommaso, and S. Dupont, Taraxacum officinale (Weber), dandelion (Asteraceae), in: Biological Control Programmes in Canada 1981-2000, edited by P. G. Mason and J. T. Huber (CABI Publishing, Wallingford, Oxon, UK, 2002) pp. 427-430.

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31. M. H. Abu-Dieyeh, J. Bernier, and A. K Watson, Sclerotinia minor advances fruiting and reduces germination in dandelion (Taraxacum officinale). Biocontrol Sci. Tech. 15, 815-825 (2005).

32. M. H. Abu-Dieyeh and A. K. Watson, Effect of turfgrass mowing height on biocontrol of dandelion with Sclerotinia minor. Biocontrol Sci. Tech. 16, 509-524 (2006).

33. M. H. Abu-Dieyeh and A. K Watson, Suppression of Taraxacum officinale populations by Sclerotinia minor and grass over-seeding. J. App. Ecol. (in press) (2006).

34. M. H. Abu-Dieyeh and A. K Watson. Efficacy of Sclerotinia minor for dandelion control: effect of dandelion accession, age and grass competition. Weed Res. (in press) (2006).

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STEVE HALLETT Education: 1987-1991 Ph.D. Biological Control of Weeds Lancaster University, UK 1984-1987 B.Sc. Biology Lancaster University, UK Academic Appointments: 2006-pres. Associate Professor, Department of Botany & Plant Pathology, Purdue University 2001-2006 Assistant Professor, Department of Botany & Plant Pathology, Purdue University 1998-2000 Lecturer, School of Agriculture & Horticulture, University of Queensland, Australia 1993-1995 Assistant Professor, Department of Plant Science, McGill University, Québec, Canada Awards and Honors:

Outstanding Undergraduate Teaching Award, Department of Botany & Plant Pathology, 2009, 2010,

2011, 2012, 2013. Outstanding Undergraduate Counselor Award, Department of Botany & Plant Pathology, 2008, 2010. Award of Excellence, Weed Science Society of America, 2001. "Seeds for Success" Acorn Award for significant grants to Purdue University. 2012. (Co-PI of Organic

Research Grant >$1,000,000) Dr. Hallett’s research has investigated the ecology of plant pathogen interactions including the development of bioherbicides for weed control, the ecological mechanisms of herbicide resistance and the ecology of weed-soil microbial community interactions in natural and agricultural systems. Dr. Hallett’s diverse research program employs a wide range of different techniques and approaches, from conventional quantitative empirical laboratory and field studies, to qualitative human dimensions surveys, and phenomenological modeling. His research addresses some of the grand challenges of agriculture in the coming decades with clearly and strategically designed research designed to lead toward the development of sustainable crop and pest management systems. Dr Hallett’s group has performed a range of studies with the Amaranthus-Microspaheropsis amaranthi pathosystem with a view to developing a bioherbicide product for the control of weedy amaranths in midwestern crop production systems. A sequence of papers has been published examining the variable responses of common waterhemp to the herbicide glyphosate, the climatic constraints on M. amaranthi, optimal spray application parameters, the interactions between M. amaranthi and chemical herbicides and the field efficacy of the candidate bioherbicide. This research has been performed in Indiana and Illinois in collaboration with Prof. Gordon Rosskamp (Western Illinois Univ.) and Dr. Loretta Ortiz-Ribbing (Univ. Illinois, Urbana Champaign). Dr. Hallett’s research with this system has developed a detailed understanding of the key opportunities and limitations for the development of a host restricted bioherbicide into a major field crop. Dr Hallett has been involved in bioherbicides research throughout his career and has published over twenty refereed journal articles in the field. His research, performed on four different continents, has consistently challenged existing theories and developed novel concepts. He is the author of the most recent comprehensive review of the subject. Dr. Hallett was involved in research on the Sarritor (Sclerotinia minor) turf bioherbicide project at McGill University, under the guidance of Dr. Alan Watson from 1991-1995.

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SELECTED PUBLICATIONS (Recent or related) i) Books Hallett, SG. 2013. The Efficiency Trap: Finding a Better Way to Achieve a Sustainable Energy Future.

Prometheus Books, Amherst, NY. 385 pp. Hallett, SG & JS Wright. 2011. Life without Oil: Why We Must Shift to a New Energy Future.

Prometheus Books, Amherst, NY. 435 pp. ii) Refereed Journal Articles: Schafer, JR, SG Hallett & WG Johnson. 2013. Soil microbial root colonization of glyphosate-treated

giant ragweed (Ambrosia trifida), horseweed (Conyza canadensis), and common lambsquarters (Chenopodium album) biotypes. Weed Science 61: 289-295.

Schafer, JR, SG Hallett & WG Johnson. 2013. Response of giant ragweed (Ambrosia trifida), horseweed

(Conyza canadensis), and common lambsquarters (Chenopodium album) biotypes to glyphosate in the presence and absence of soil microorganisms . Weed Science 60: 641-649.

Ortiz-Ribbing, LM., KR Glassman, GK Roskamp & SG Hallett. 2011. Performance of two bioherbicide

fungi for waterhemp and pigweed control in pumpkin and soybean. Plant Dis. 95:469-477. Shabana, Y, D Singh, LM Ortiz-Ribbing & SG Hallett. 2010. Production and formulation of high quality

conidia of Microsphaeropsis amaranthi for the biological control of weedy Amaranthus species. Biol. Contr. 55:49-57.

Stewart, JM, Latin, R, Reicher, Z, and Hallett, SG. 2008. Influence of trinexapac ethyl on the efficacy

of chlorothalonil and propiconazole for control of dollar spot on creeping bentgrass. Online. Applied Turf Science. doi: 10.1094/ATS-2008-0319-01-RS.

Callaway, RM, D Cipollini, K Barto, GC Thelen, SG Hallett, D Prati, K Stinson & JN Klironomos.

2008. An invasive plant suppresses fungal mutualisms in America but not in its native Europe. Ecology 89:1043-1055.

Beed, FD, SG Hallett, J Venne & AK Watson. 2007. Biocontrol using Fusarium oxysporum: a critical

component of integrated Striga management. In: J Gressel & G Ejeta (eds) Integrating New Technologies for Striga Control: Towards Ending the Witch-Hunt. World Scientific Publ. Co. Inc., Hackensack, NJ.

Davis, AS, KI Anderson, SG Hallett and KA Renner. 2006. Weed seed mortality in soils with

contrasting agricultural management histories. Weed Sci. 54:291-297. Hallett, SG. 2006. Dislocation from coevolved relationships: a unifying theory for plant invasion and

naturalization? Weed Sci. 54:282-290. Smith, DA, DA Doll, D Singh & SG Hallett. 2006. Climatic constraints to the potential of

Microsphaeropsis amaranthi as a bioherbicide for common waterhemp. Phytopathology 96:308-312.

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Smith, DA & SG Hallett. 2006. Interactions between chemical herbicides and the candidate bioherbicide Microsphaeropsis amaranthi. Weed Sci. 54:532-537.

Smith, DA & SG Hallett. 2006. Variable response of common waterhemp (Amaranthus rudis Sauer) to

glyphosate. Weed Technol. 20:466-471. Stinson, KA, S Campbell, JR Powell, BE Wolfe, RM Callaway, GC Thelen, SG Hallett, D Prati & JN

Klironomos. 2006. Invasive plant suppresses the establishment and growth of native trees by allelochemical disruption of belowground mutualists. PLoS Biology 4:727-731.

Doll, DA, PE Sojka & SG Hallett. 2005. Factors affecting the efficacy of spray applications of the

bioherbicidal fungus Microsphaeropsis amaranthi. Weed Technol. 19:110-115. Brière, SC, AK Watson, TC Paulitz & SG Hallett. 2000. Oxalic acid production and mycelial biomass

yield of Sclerotinia minor for the formulation enhancement of a granular turf bioherbicide. Biocontrol Sci. Technol. 10: 281-289.

Brière, SC, AK Watson, TC Paulitz & SG Hallett. 1995. First report of a Phoma sp. on common

ragweed in North America. Plant Dis. 79:968.

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Last updated 3/28/2012

Aaron J. Patton Abbreviated Curriculum Vitae Purdue University Department of Agronomy 915 West State Street West Lafayette, IN 47907-2054 Cellular Phone (765) 414-5131 Office Phone: (765) 494-9737 Fax: (765) 296-6335 Email: [email protected] Website: http://www.agry.purdue.edu/turf Education Ph.D. in Agronomy, Purdue University, West Lafayette, IN. December 2006. M.S. in Agronomy, Purdue University, West Lafayette, IN. August 2003. B.S., in Horticulture, Iowa State University, Ames, IA. December 2000. Professional Experience Associate Professor/Turfgrass Extension Specialist, Purdue University, Department of Agronomy, July 2010 to current (60% Extension, 20% Research, 20% Teaching) Dr. Patton’s primary responsibility is to develop and deliver educational programs to the turfgrass industry of Indiana and the Midwest. Therefore, he is also actively involved in applied research and helps to maintain a high quality undergraduate program. The core Turfgrass Science Program includes 6 faculty, 4 professional staff, 5-10 graduate students and 1 clerical staff. In addition, up to 4 more faculty and several upper level AP staff work closely with the program. As part of the Purdue Turf Team, Dr. Patton fills a role as a leader of the applied research and Extension/outreach efforts. Dr. Patton is Director of the W.H. Daniel Turfgrass Research and Diagnostic Center. Dr. Patton supervises the Center’s manager, coordinates external funding and donations for the Center and is responsible for long-term strategic planning for the facility and repair and rehabilitation funds. The Daniel Center is used by approximately 4,000 people annually including programs by academic departments, intercollegiate athletics, and industry in addition to the Extension and teaching activities of the Turf Team. 2013-current Associate Professor/ Turfgrass Extension Specialist, Purdue Univ., Dept. of Agronomy. 2010-2013 Assistant Professor/ Turfgrass Extension Specialist, Purdue Univ., Dept. of Agronomy. 2006-2010 Asst. Professor/ Turfgrass Extension Specialist, Univ. of Arkansas, Dept. of Horticulture Research Author or junior author of 35 refereed publications, 4 refereed proceedings, 51 abstracts, 7 research reports, 114 experiment station research articles, and 127 technical reports for funding agencies. Responsible for over $1.1 M in research support since becoming a faculty member in 2006.

Relevant Refereed Publications (last 3 years shown) 1. Beck, L.L., A.J. Patton*, and D.V. Weisenberger. 2014. Mowing before or after an herbicide

application does not influence ground ivy (Glechoma hederacea) control. Appl. Turfgrass Sci. In press.

2. Sousek, M.D., R.E. Gaussoin, A.J. Patton, D.V. Weisenberger, and Z.J. Reicher. 2013. Weed control and turf safety of single and sequential applications of herbicides over spring seedings. Appl. Turfgrass Sci. In press.

3. Patton, A.J., G.E. Ruhl, T.C. Creswell, P. Wan, D. Scott, J. Becovitz, and D.V. Weisenberger. 2013. Potential damage to sensitive landscape plants from wood chips of aminocyclopyrachlor damaged trees. Weed Technol. In press.

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4. Brosnan, J.T., G.K. Breeden, A.J. Patton, and D.V. Weisenberger. 2013. Triclopyr reduces smooth crabgrass bleaching with topramezone. Appl. Turfgrass Sci. doi:10.1094/ATS-2013-0038-BR.

5. Elmore, M.T., J.T. Brosnan, G.K. Breeden, and A.J. Patton. 2013. Mesotrione, topramezone and amicarbazone combinations for postemergence annual bluegrass (Poa annua) control. Weed Technol. 27:596-603.

6. Patton, A.J., D.V. Weisenberger, J.T. Brosnan, and G.K. Breeden. 2013. Safety of labeled herbicides for broadleaf weed control in creeping bentgrass putting greens. Turfgrass Sci. doi:10.1094/ATS-2013-0523-01-BR.

7. Brosnan, J.T., G.K. Breeden, M.T. Elmore, A.J. Patton, and D.V. Weisenberger. 2012. Zoysiagrass seedhead suppression with imidazolinone herbicides. Weed Technol. 26:708-713.

8. Proctor, C.A., M.D. Sousek, A.J. Patton, D.V. Weisenberger, and Z.J. Reicher. 2012. Combining preemergence herbicides in tank-mixes or as sequential applications provides season-long crabgrass control. HortScience 47(8):1159-1162.

9. Jellicorse, W.R., M.D. Richardson, J.H. McCalla, D.E. Karcher, A.J. Patton, and J.W. Boyd. 2012. Seeded bermudagrass establishment in an overseeded perennial ryegrass stand as affected by transition herbicide and seeding date. Appl. Turfgrass Sci. doi:10.1094/ATS-2012-0721-01-RS.

10. Reicher, Z.J., A.J. Patton, and D.V. Weisenberger. 2012. Suppression of field paspalum in Kentucky bluegrass with mesotrione. [Online]. Appl. Turfgrass Sci. doi:10.1094/ATS-2012-0626-01-RS.

Teaching Dr. Patton integrates his applied research program and his experience working with clientele in his Extension program to prepare undergraduate students for real-life challenges in the turfgrass capstone course he teaches each fall. He also teaches a Professional Presentations course for graduate students in which he is able to take his passion for mentoring students and use it to prepare them for future success. Dr. Patton has Taught, co-taught, or guest lectured in 16 courses at three Universities with an average instructor rating of 4.7/5.0. He developed two courses and instructional materials for 5 separate courses. Published 1 book chapter on turfgrass management for use in classroom instruction and an abstract on mentoring TA’s. Extension Author or junior author of 51 peer reviewed extension publications and 25 trade or popular press articles. Author of 1 book chapter, 3 refereed scholarship of extension manuscripts and 5 abstracts. Developed website with over 100,000 page views annually, maintain a turf tips blog (listserv) with over 2,600 subscribers and over 50 posts annually, and spoke at over 300 state, regional, or national meetings to over 20,000 people since 2001.

Honors and Awards Faculty

Individual Awards Purdue Cooperative Extension Specialists' Association, Special Award for Program

Enhancement, 2013 Dept. of Agronomy Nominee for the R.L. Kohls Early Career Award, College of Ag., Purdue

University, 2012 Awardee, Purdue University College of Agriculture, Service Learning Faculty Development

Program, 2012 Extension Excellence Award for Early Career-Specialist, University of Arkansas Division of

Agriculture, 2009

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Team Awards Team Award, Purdue University College of Agriculture, Imprelis Herbicide Injury Response

Team, 2013 Award of Achievement, Midwest Regional Turf Foundation, Imprelis Response Team, 2013 Team Award, Purdue University Cooperative Extension Specialists Association, Imprelis

Herbicide Injury Response Team, 2012 Service and Leadership Activities (last three years)

National Education Committee, Golf Course Superintendents Association of America, 2013-current Secretary, NCERA221 Program, 2012-2014, Chair in 2013 and 2014 Senior Editor, Applied Turfgrass Science Journal, Plant Management Network, Crop Science

Society of America (C311), 2013-2015; Fred V. Grau Turfgrass Science Award Committee (C458). Crop Science Society of America,

2013-2015; Chair in 2014 Early Career Representative, American Society of Agronomy Board of Directors (A003), 2011-

2013 Science Policy Committee, American Society of Agronomy (A537), 2011-2013 Early Career Members Committee (ACS530), American Society of Agronomy, 2010-2013 C-5 Division (Turfgrass) symposium committee, Crop Science Society of America, 2010-2013;

Chair in 2010-2011. C-5 Division (Turfgrass) Extension symposium committee, Crop Science Society of America,

2007-2013; Chair in 2007-2008 Acquisitions Editor, Applied Turfgrass Science Journal, Plant Management Network, Crop

Science Society of America (C311), 2008-2011 Reviewer: Agronomy Journal, Amer. J. of Hort. Sci., Applied Turfgrass Science, Crop Science,

Crop Science Monograph, HortScience, HortTechnology, the International Turfgrass Research Society Journal, Journal of Environmental Quality, Weed Science, and Weed Technology. Total of 38 manuscripts reviewed since 2006.

Service on State and/or Governmental Committees Executive Director and education chair of the Midwest Regional Turf Foundation (2010-present) Pesticide Training – Category 3b Advisory Committee for the OISC (2010-present) Indiana Golf Course Owners Association Advisory Committee (2011-present) Departmental Committee and Service Department Head Advisory Committee, Department of Agronomy (2011-present). Department Seminar Committee, Department of Agronomy, (co-chair, 2013-present). Distinguished Alumni, Honorary Degree, and Outstanding Dept. Alumni Award Committee

(2012-present)

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- Biographical Sketch pg. 1 of 3 -

Jenny T. Kao-Kniffin Department of Horticulture Cornell University office: (607) 255-8886 134a Plant Sciences Building fax: (607) 255-0599 Ithaca, NY 14853-5904 E-mail: [email protected]

A. PROFESSIONAL PREPARATION

State University of New York at Binghamton Environmental Studies B.S. 1999 State University of New York at Binghamton Biological Sciences M.S. 2002 University of Wisconsin-Madison Land Resources Ph.D. 2007

B. APPOINTMENTS 2010-present Assistant Professor, Cornell University, Ithaca, NY, Department of Horticulture 2009-2010 NSF Postdoctoral Research Fellow, Office of Polar Programs, field site: Barrow, AK 2008 Visiting Scientist, National Ecological Observatory Network (NEON), Boulder, CO

C. 5 PUBLICATIONS MOST CLOSELY RELATED TO THE PROJECT

Kao-Kniffin, J. and T.C. Balser. 2010. Soil microbial composition and nitrogen cycling in a disturbed wet prairie restoration (Wisconsin). Ecological Restoration 28:20-22 DOI:10.3368/er.28.1.20

Zhu, B., K. Panke-Buisse, and J. Kao-Kniffin. In review. Nitrogen fertilization shows minimal influence on plant rhizosphere effects

Kao, J.T., J.E. Titus, and W. Zhu. 2003. Differential nitrogen and phosphorus retention by five wetland plant species. Wetlands 23:979-987 DOI: 10.1672/0277-5212(2003)023[0979:DNAPRB]2.0.CO;2

Kao-Kniffin, J. and T.C. Balser. 2008. Soil fertility and the impact of exotic invasion on microbial communities in Hawaiian forests. Microbial Ecology 56:55-63 DOI 10.1007/s00248-007-9323-1

Kao-Kniffin, J. and B, Zhu. 2013. A microbial link between elevated CO2 and methane emissions that is plant species-specific. Microbial Ecology 66:621-629 DOI: 10.1007/s00248-013-0254-8

5 OTHER SIGNIFICANT PUBLICATIONS Kao-Kniffin, J., S.M. Carver, and A. DiTommaso. 2013. Advancing weed management strategies using

metagenomic techniques. Weed Science 61:171-184 DOI: 10.1614/WS-D-12-00114.1

Kao- Kniffin and T.C. Balser. 2007. Elevated CO2 differentially alters belowground plant and soil microbial community structure in reed canary grass-invaded experimental wetlands. Soil Biology & Biochemistry 39:517-525 DOI:10.1016/j.soilbio.2006.08.024

Kao- Kniffin, J., D. Freyre, and T.C. Balser. 2011. Increased methane emissions from an invasive wetland plant under elevated carbon dioxide levels. Applied Soil Ecology 48:309-312 DOI: 10.1016/j.apsoil.2011.04.008

Kao-Kniffin, J. 2012. Rhizosphere Ecology. McGraw Hill Encyclopedia of Science & Technology, 11th Edition

Kao-Kniffin, J., D. Freyre, and T.C. Balser. 2010. Methane dynamics across wetland plant species.

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Aquatic Botany 93:107-113 DOI:10.1016/j.aquabot.2010.03.009 5 EXTENSION PUBLICATIONS Kao-Kniffin, J. 2012. Regulatory Restrictions on Pesticides in Turf. NYSTA Online Newsletter. August issue.

Kao-Kniffin, J. 2011. Weed management under the school pesticide ban. Cornell University Turfgrass Times (CUTT) 22(2):12.

Kao-Kniffin, J. 2011. Allowable herbicides Child Safe Playing Fields Law. Cornell University Turfgrass Times (ShortCUTT) 26:1-2.

Kao-Kniffin, J., F. Rossi., and L. Weston. 2013. Pest Management Guidelines for Commercial Turfgrass, Weed Management Chapter 7. Cornell Cooperative Extension Publication.

Kao-Kniffin, J., 2013. Weed management on school grounds in New York State. Cornell Turfgrass publication.

D. SYNERGISTIC ACTIVITIES

D.1. Service Activities: Member, NSF IGERT Cross-Scale Biogeochemistry and Climate, Cornell University (2012-present) Faculty Fellow, Flora Rose Hall Residence, Cornell University, 2012-present Faculty Fellow, Atkinson Center for a Sustainable Future, Cornell University, 2011-present Invited Participant, Emerging Frontiers in Rhizosphere Science (NSF-sponsored workshop), Airlie Center, Warrenton, VA (2011) Grant Reviewer for: University of California ANR Grants Program (2012); Toward Sustainability Foundation Grants Program (2010-present); U.S. Department of Energy’s (DOE) Office of Science, Office of Biological and Environmental Research (2013) Journal Reviewer for: Applied Soil Ecology, Wetlands, Agronomy, Environmental Engineering, Journal of Environmental Quality, Sustainability Science, Wetlands Ecology & Management, BMC Bioinformatics

D.2. Professional Development: Received advanced training at the following research centers and universities: 1. (2011) Technische Universität München (TUM) training on soil organic matter techniques involving NanoSIMS, 13C NMR, soil fractionation, and confocal microscopy; 2. (2009) Cold Spring Harbor Laboratory (CSHL) training program on laboratory and bioinformatic analysis of genomic datasets produced using next-generation sequencing techniques (Illumina, 454 Roche, and ABI Solid); 3. (2008 and 2009) Biopharmaceutical Technology Center Institute (BTCI) training programs on statistical and bioinformatic approaches to analyzing large genomic microarray datasets and new applications of RT-PCR; 4. (2007) National Center for Atmospheric Research (NCAR) Advanced Study Colloquium on Regional Biogeochemistry program that focused on finding and discussing new research and methodologies using top-down and bottom-up approaches in regional biogeochemistry; 5. (2006) NSF Advance VT, Transforming the Professoriate Conference (Virginia Tech) training program focused on providing women and minorities with the tools to plan for a successful career in the sciences and academia.

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D.3. Educational Activities: Teaching Fellow: (summer 2006) Howard Hughes Medical Institute (HHMI) Teaching Fellows Program Intern: (2005) Delta Program, Center for the Integration of Research, Teaching & Learning (CIRTL), University of Wisconsin-Madison); NSF-sponsored program

Invited Panelist: Future Faculty Professional Development: Effectively engaging graduate students as co-investigators of student learning, 2006 Carnegie Academy for the Scholarship of Teaching and Learning (CASTL) Colloquium on the Scholarship of Teaching and Learning, Madison, WI

E. COLLABORATORS & OTHER AFFILIATIONS E.1. Collaborators and Co-Editors: Nina Bassuk (Cornell U.) James Bockheim (UW-Madison),

Sarah Carver (Cornell U.), Antonio DiTommaso (Cornell U.), Laurie Drinkwater (Cornell U.), Peter Groffman (Cary Institute), Jo Handelsman (Yale U.), Carsten Mueller (Technische Universität München), Biao Zhu (Berkeley National Laboratory)

E.2. Graduate Advisors and Postdoctoral Sponsors: Teri C. Balser (U of Florida, Soil and Water

Dept.), Jo Handelsman (Yale University, Molecular, Cellular, and Developmental Biology Dept.), James Bockheim (UW-Madison, Soil Science Dept.), John Titus (S.U.N.Y. Binghamton, Biology Dept.)

E.3. Ph.D. Committee Advisors: Monica G. Turner (UW-Madison, Zoology Dept.), Jo Handelsman

(Yale University, Molecular, Cellular, and Developmental Biology Dept.), Joy B. Zedler (UW-Madison, Botany Dept.), Art McEvoy (UW-Madison, Law School)

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A. DiTommaso

ANTONIO DITOMMASO Department of Crop & Soil Sciences Phone: (607) 254-4702 903 Bradfield Hall Fax: (607) 255-2644 Cornell University E-mail: [email protected] Ithaca, New York 14853 USA EDUCATION 1995 Ph.D., Weed Ecology, McGill University, Montreal, Quebec, Canada. 1989 M.S., Plant Ecology, Queen’s University, Kingston, Ontario, Canada. 1986 B.S.(Agr.), Environmental Biology, McGill University, Montreal, Quebec, Canada. PROFESSIONAL EXPERIENCE July 2005 - Associate Professor, Department of Crop & Soil Sciences, Cornell University 1999-2005 Assistant Professor, Department of Crop & Soil Sciences, Cornell University 1997-1999 Faculty Lecturer, Department of Plant Science, McGill University 1995-1997 Assistant Professor (Special category), Dept. of Plant Science, McGill University 1992 Sessional Lecturer, Department of Plant Science, McGill University AREAS OF EXPERTISE Primary areas of scholarship focus on:

1. Effects of the environment on weed species. Evaluation of the effects of biotic (e.g. plant competition; selective disease and insect predation) and abiotic factors (e.g. light, fertility, salinity) on the seed biology, growth and reproduction of important agricultural weeds (common ragweed, velvetleaf, pigweeds) and introduced invasive plant species (e.g. swallow-wort, mugwort) of natural communities in the Northeastern United States.

2. Manipulation of the environment to suppress weeds. Research aimed at modifying the biotic and/or abiotic environment of a target weed to manage troublesome weeds in both cropland and natural systems. These tactics can be used either alone or as part of an integrated management strategy. Manipulation of the biotic environment is primarily achieved through the use of selective biological control agents such as fungal pathogens and insects while abiotic manipulations focus largely on soil fertility management.

MEMBERSHIP IN PROFESSIONAL SOCIETIES American Society of Agronomy and Crop Science Botanical Society of America Canadian Weed Science Society European Weed Research Society International Biocontrol Group International Weed Science Society Northeastern Weed Science Society Quebec Society for the Protection of Plants Weed Science Society of America EFFORT DISTRIBUTION Research: 45% Teaching: 55% PROFESSIONAL ACTIVITIES Past-President, Northeastern Weed Science Society [2013– 2014]

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A. DiTommaso

Associate Editor, Invasive Plant Science and Management [2013– present] Associate Editor, Weed Technology [2007 – 2013] Member of Publications Board, Weed Science Society of America (WSSA) [2009-2013] Member of the Outstanding Researcher Award Committee of the WSSA [2011-2014] REFEREED PUBLICATIONS (2011-present) DiTommaso, A., Q. Zhong, and D.R. Clements. 2014. Identifying climate change as a factor in the establishment and persistence of invasive weeds in agricultural crops. In: L.H. Ziska and J.S. Dukes, eds. Invasive Species and Climate Change. CABI Publishing,Wallingford, UK. pp. 000-000. In Press. DiTommaso, A., L.R. Milbrath, T. Bittner, and F.R. Wesley. Pale swallowwort (Vincetoxicum rossicum) response to cutting and herbicides. 2013. Invasive Plant Science and Management 6(3):381-390. Kao-Kniffin, J., S. M. Carver, and A. DiTommaso. 2013. Development of metagenomic tools to advance weed management strategies. (Invited paper) Weed Science. 61(2)171-184. Magidow, L.C., A. DiTommaso, Q.M. Ketterings, C.L. Mohler, and L.R. Milbrath. 2013. Emergence and performance of two invasive swallow-worts (Vincetoxicum spp.) in contrasting soil types and soil pH. Invasive Plant Science and Management. 6(2):281-291. Eom, S.H., A. DiTommaso, and L.A. Weston. 2013. Effects of soil salinity in the growth of Ambrosia artemisiifolia biotypes collected from roadside and agricultural field. Journal of Plant Nutrition. 36(14) 2191-2204. Goulet, E.J., J. Thaler, A. DiTommaso, M. Schwarzlander, and E.J. Shields. 2013. Impact of Mecinus janthinus (Coleoptera: Curculionidae) on the growth and reproduction of Linaria dalmatica. The Great Lakes Entomologist. 46(1-2):90-98. Zhong, Q.., D-J Mao, G-M Quan, J-E Zhang, J-F Xie, and A. DiTommaso. 2013. Antioxidant response of the invasive herb Ambrosia artemisiifolia L. to different irradiance intensities. Phytoprotection. 93(1): 8-15. Bravo, M., A. DiTommaso, and D. Hayes. 2012. Exotic plant inventory, landscape survey and invasiveness assessment: Roosevelt-Vanderbilt National Historic Sites, Hyde Park, NY. HortTechnology 22(5):682-693.

Clements, D.R. and A. DiTommaso. 2012. Predicting weed invasion in Canada under climate change: measuring evolutionary potential. Canadian Journal of Plant Science 92(6):1013-1020.

Darbyshire, S. J., A., Francis, A. DiTommaso, and D. R. Clements. 2012. The Biology of Canadian Weeds. 150. Erechtites hieraciifolius (L.) Raf. ex DC. Canadian Journal of Plant Science 92(4):729-746.

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A. DiTommaso

Mohler, C.L., C. Dykeman, E.B. Nelson, and A. DiTommaso. 2012. Reduction in weed seedling emergence by pathogens following the incorporation of green crop residue. Weed Research 52(5): 467-477.

Orlowski, J., W.J. Cox, A. DiTommaso, and W. Knoblauch. 2012. Planting soybean with a grain drill inconsistently increases yield and profit. Agronomy Journal 104:1065-1073.

Smith, E.A., A. DiTommaso, M. Fuchs, A.M. Shelton, and B.A. Nault. 2012. Abundance of weed hosts as potential sources of onion and potato viruses in western New York. Crop Protection 37:91-96.

Stephens, E.J., J.E. Losey, L.L. Allee, A. DiTommaso, C. Bodner, and A. Breyre. 2012. The impact of Cry3Bb1 Bt-maize on two guilds of beneficial beetles. Agriculture, Ecosystems and Environment 156:72-81.

Zhong Q., Dan Juan, M., Guo, M-Q, Jia-en, Z. Jun, F-X, and A. DiTommaso. 2012. Physiological and morphological responses of invasive Ambrosia artemisiifolia (common ragweed) to different irradiances. Botany 90(12):1284-1294. Averill, K.M., A. DiTommaso, C.L. Mohler and L.R. Milbrath. 2011. Survival, growth, and fecundity of the invasive swallow-worts (Vincetoxicum rossicum and V. nigrum) in New York State. Invasive Plant Science and Management 4(2):198-206. Burke, J.M. and A. DiTommaso. 2011. Antigonon leptopus (Corallita): Intentional introduction of a plant with documented invasive capability. Invasive Plant Science and Management 4(3):265-273. Clements, D.R. and A. DiTommaso. 2011. Climate change and weed adaptation: Can evolution of invasive plants lead to greater range expansion than forecasted? Weed Research 51(3): 227-240.

DiTommaso, A. 2011. Pale Swallow-wort: An Emerging Threat to Natural and Seminatural Habitats in the Lower Great Lakes Basin of North America. In: Restoration Ecology. Greipsson, S. [ed.], Jones and Bartlett Learning, Sudbury, MA. pp.169-174.

Francis, A., S.J. Darbyshire, D.R. Clements, and A. DiTommaso. 2011. The Biology of

Canadian Weeds. 146. Lapsana communis L.. Canadian Journal of Plant Science 91(3):553-569. Smith, E.A., A. DiTommaso, M. Fuchs, A.M. Shelton, and B.A. Nault. 2011. Weed hosts for

onion thrips (Thysanoptera: Thripidae) and their potential role in the epidemiology of Iris yellow spot virus in an onion ecosystem. Environmental Entomology 40(2):194-203.

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Biographical Sketch

Joseph Crowell Neal

Affiliation: Department of Horticultural Science

North Carolina State University

Raleigh, NC 27695

Education Ph.D., 1984 North Carolina State University, Raleigh, NC, Horticultural Science,

M.S., 1981 Clemson University, Clemson, SC, Horticulture

B.S., 1978 University of Georgia, Athens, GA, Horticulture

Previous Professional Experience 11/84 to 9/96: Assistant and Associate Professor of Weed Science, Department of Floriculture

and Ornamental Horticulture, Cornell University

9/96 to present: Professor and Extension Specialist, Department of Horticultural Science, North

Carolina State University

Current Professional Responsibilities:

Responsibilities include: research and extension education for weed management systems in

nursery crops, urban landscapes and Christmas trees. Research interests include integrated weed

management, biological control of weeds, biology and management of invasive weeds. Outreach

education programs emphasize curriculum-based educational programs.

Scholarly and Professional Honors 1990 Tools in Teaching award from Epsilon Sigma Phi, Lambda Chapter - The Cooperative

Extension Honorary Society.

1993 Northeastern Weed Science Society Award for "Outstanding Applied Research in Turf,

Ornamentals, or Conservation Acreage".

1997 Outstanding Extension Publication Award from Epsilon Sigma Phi, Lambda Chapter

1998 Educational Materials Award –American Society for Horticultural Science

2004 Fellow – Northeastern Weed Science Society

2006 Distinguished Service Award – Amer. Soc. for Horticultural Sci, Nursery Working Group

2006 Outstanding Extension Publication Award – Amer. Soc. for Hort Sci, Extension Division

2008 Porter Henegar Award -- Southern Nursery Association

2009 Outstanding Researcher Award -- Northeastern Weed Science Society

2009 Educational Materials Award -- American Society for Horticultural Science

2013 Kim Powell Award for service to the industry – NC Nursery and Landscape Assoc.

Professional Society Memberships & Service:

- American Society for Horticultural Science

- International Bioherbicide Group (IBG)-- IXth IBG Workshop organizing committee chair 2009

- Northeastern Weed Science Society

Graduate student presentation judging committee, 1998-2004; Nominating committee, 1989,

1991, 1992 (chair); Chair, resolutions committee, 1990; Executive committee, 1993-1998;

Editor, 1993 – 1994; Vice President and program chair, 1995; President-Elect, 1996;

President, 1997; Awards committee, 1998-2003; Chair, biological control section, 2000

- North Carolina Exotic Pest Plant Council

Organizing committee 2000-2001; Nominating committee 2001; Vice-President 2004

- Southern Nurserymen’s Association Research Conference

Student presentation judge, 1998-1999 (chair); Chair weed control section, 2003-2004

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Biographical Sketch

- Weed Science Society of America

Chair, turf and ornamentals section, 1990; Chair, biocontrol section, 2000; Chair, symposium

committee: Fate of Herbicides Applied to Turf and Ornamentals, 1992; Awards committee 2003-

2006; Board of Directors 2006-2010; Chair Biocontrol committee 2006-2008.

Related Grants 1 Field-faculty pest management in-service training; 1999, 2001, 2002, 2004, NCSU IPM grants

$8000

2. Herbicide safety in ornamentals; annual grants from USDA IR-4 $7000 to $18,000 per year

3. Development of a prototype digital weed identification tool (Weed IT) for use online or in the

field via personal digital assistants (PDAs) or smartphones. (w/ A. Krings) NCSU Univ. Outreach

and Engagement Grant $10,000; SR-IPM enhancement grant $25,000 (2008-2012)

http://www.cals.ncsu.edu/plantbiology/ncsc/containerWeeds/

4. Development of an invasive assessment protocol for NC. NC Nursery and Landscape

Association $36,000 (2008-2009)

5. Bio-based control of broadleaf weeds in turf and landscapes. USDA-IR-4 BioPesticide Program,

Marrone BioSciences, Neudorff Co, and The Scotts Co: $70,000

Publications Summary

Refereed Journal Articles: 50; Reviewed Proceedings Papers: 6; Abstracts of Presentations at

Scientific Conferences: 169; Books: 1 (Weeds of the Northeast); Book Chapters: 4; Reviewed and edited

Extension Bulletins: 8; Extension newsletter articles and fact sheets: 40; Popular press: 24

Graduate Students:

Chaired: 4 Ph.D., 9 M.S.

Advisory Committee: 5 Ph.D.; 7 M.S.

Selected Publications:

Fulcher, A;. Klingeman; Juang-Horng Chong; LeBude;. Armel; Chappell; Frank;. Hale; Neal; White;

Williams-Woodward; Ivors; Adkins; Senesac; Windham. 2012. Stakeholder Vision of Future

Direction and Strategies for Southeastern U.S. Nursery Pest Research and Extension Programming.

Journal of Integrated Pest Management 3(2):D1-D8. DOI: http://dx.doi.org/10.1603/IPM11030

Post, A. R., R. Ali, A. Krings, Qui-Yun (Jenny) Xiang, B. R. Sosinski and J. C. Neal. 2011. On the

identity of the weedy Cardamine (Brassicaceae) species in United States nurseries: Evidence from

molecules and morphology. Weed Science WeedScience 59:123-135

Walker, L.H., J.C. Neal and J.Derr. 2010. Preemergence control of doveweed (Murdannia nudiflora) in

container grown nursery crops. J. Environ. Hort. 28(1):8-12

LeBude, A., B. L. Upchurch, and J. C. Neal. 2009. Preemergence Herbicide Applications to Six

Containerized Woody Ornamental Rootstocks do not Affect Winter Grafting Success. J. Environ.

Hort. 27(2):119-122.

LeBude, A., T. Bilderback, J. Neal, and C. Safeley. 2008. Best Management Practices for Field

Nurseries. NC Assoc. of Nurserymen, Raleigh.

Neal, J. C. and J. F. Derr. 2005. Weeds of container nurseries in the United States. NC Nurserymen’s

Assoc. 16 p

Judge,C. A. and J. C. Neal. 2006. Early postemergence control of nursery weeds with Broadstar, OH2

and Snapshot TG. J. Environ. Hort. 24(2):105-108

Penny, G.M. and J. C. Neal. 2003. Germination of mulberry weed (Fatoua villosa) as affected by light,

planting depth, mulching and temperature. Weed Technology 17:213-218.

Uva, R. H., J. C. Neal and J. M. DiTomaso. 1997. Weeds of the Northeast. Cornell University Press,

New York. 396pp.

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http://www.cals.ncsu.edu/plantbiology/ncsc/containerWeeds/

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Documents

CRITERIA FOR RANKING EVALUATIONS OF IR-4 ADVANCED …ir4.rutgers.edu/Biopesticides/Proposals/15A.pdf · 2. Experimental design, work plan and preliminary research. of 10 3. Does experimental