Sustainable Management of Soil-borne Diseases in Nursery Production

Final report for GS16-155

Project Type: Graduate Student
Funds awarded in 2016: $11,000.00
Projected End Date: 08/31/2018
Grant Recipient: Tennessee State University
Region: Southern
State: Tennessee
Graduate Student:
Major Professor:
Dr. Fulya Baysal-Gurel
Tennessee State University
Expand All

Project Information

Summary:

Soilborne diseases are the most serious production problem of Southern region nursery producers and these producers have cited better, more effective alternative soilborne disease management as their highest priority need. The goal of the proposed research is to improve soilborne disease management and nursery production efficiency through effective applications of biofumigant cover crops in nursery production. The use of biofumigant cover crops has been explored most extensively in vegetable, fruit and flower production. To date, their use and value have not been documented in woody ornamental nursery production. Specific questions to be addressed in this research include: 1) What are the effects of using biofumigant cover crops (oilseed radish, mustard (white, Mighty Mustard® Pacific Gold, oriental, red giant and Amara) turnips (purple top forage), arugula (Sylvetta Green, Wasabi, Bellezia, Dragon’s Tongue, Olive-Leaved Sylvetta and Astro) and rape (Dwarf Essex)) in nursery system in Tennessee? and 2) Which functionally-important microbial populations are most affected by biofumigant cover crops? The results of the proposed research will improve the ability of stakeholders to develop a more effective nursery production system plan. As a result of this research, nursery producers will have new options for sustainable control of soilborne diseases.

Project Objectives:

This project focuses on studying diseases of woody ornamental nursery crops with the ultimate goal of delivering sustainable soil-borne disease management strategies. Specific questions to be addressed by the proposed research include: 1) What are the effects of using biofumigant cover crops in nursery cropping system in Tennessee? and 2) Which functionally-important microbial populations (i.e. pathogens and their antagonists) are most affected by biofumigant cover crops? To answer these questions and provide nursery producers with a useful synthesis of our results the following main objective and specific objectives will be pursued:

Main objective

Assess environmentally friendly biofumigant cover crops for soilborne diseases and improved plant growth.

Specific objectives

  1. Evaluating pathogenicity of Rhizoctonia solani and Phytophthora nicotianae to Brassicaceae biofumigant cover crops

        1.1 Evaluation of inoculation methods and inoculum level of Rhizoctonia solani to determine disease response

        1.2 Pathogenicity of Rhizoctonia solani and Phytophthora nicotianae to biofumigant cover crops

  1. Determining the effect of biofumigation in suppressing Rhizoctonia solani and Phytophthora nicotianae in woody ornamental nurseries

        2.1 Evaluating for the ability of Brassicaceae cover crops to release glucosinolate hydrolyze compounds and control R. solani and P. nicotianae under greenhouse conditions

        2.2 Evaluating sustainable approaches to control soilborne diseases in woody ornamental nurseries

Research

Materials and methods:

Objective 1. Evaluating pathogenicity of Rhizoctonia solani and Phytophthora nicotianae to Brassicaceae biofumigant cover crops

Objective 1.1 Evaluation of inoculation methods and inoculum level of Rhizoctonia solani to determine disease response

Fungal culture

Rhizoctonia solani culture was obtained from the culture collection of Dr. Fulya Baysal-Gurel at Tennessee State University Otis L. Floyd Nursery Research Center McMinnville, TN. Isolate FBG201505 of R. solani, originally isolated from a diseased viburnum plant, and was grown on Potato Dextrose Agar (PDA) media in 9-cm-diameter petri dishes at 25 °C for 7 days.

Inoculation methods and inoculum levels

For this study, inoculum was prepared using chopped potato soil medium (Ko and Hora 1971; Nelson and Hoitink 1982), agar plug and agar slurry inoculation methods and each of this inoculum method was having 3 inoculum levels (Table 2.1).

Table 1. Inoculation methods and inoculum levels

Inoculation methods

Inoculum levels

Chopped potato soil medium

0.5,1.0 and 1.5 g/Kg -1mix

Agar plugs

1, 3 and 5 No. plug/ pot

Agar slurry

1, 2 and 3 petri dishes/L

 

Chopped potato soil medium inoculation method

A mixture of silt loam soil (2,000 ml), finely chopped potatoes (240 g) and sterile distilled H2O (400 ml) was autoclaved for 1 h on each of 3 consecutive days and shaken after each autoclaved. The mixture was inoculated with 7 discs (with 5 No. Plug) of R. solani grown on PDA media. The inoculated mixture was incubated at room temperature for 7 to 10 days and then air-dried for 2 to 3 days, ground with a mortar and pestle, and sieved to a 1- to 2-mm particle size. Three levels of the inoculum (0.5, 1.0 and 1.5 g kg-1 mix) was used in the experiment. Inoculum was mixed with 50 g of the sterilized soi and placed on top of the sterilized soil of the Viburnum planted pots.

Agar plug inoculation method

Agar plugs were cut from the leading edge of each actively growing Rhizoctonia colony with cork borer. Three-inoculum levels; number 1 plug/ pot, number 3 plug/ pot and number 5 plug/ pot, were used for the experiment. Each 10 cm pot filled with sterilized soil was inoculated with a single plug 1 inch below from the surface of the soil.

Agar slurry inoculation method

Seven days old Rhizoctonia culture was decanted into a sterilezed beaker with sterilezed distilled H2O (1,000 ml) and homogenized with blender (Hamilton Beach hand blender, Model number 59785R) to make slurry. Three inoculum levels; 1 petri dish/L, 2 petri dishes/L and 3 petri dishes/L were used for the experiment. Viburnum plants grown in sterilized soil were inoculated with 100 ml of prepared slurry.

Greenhouse bioassay

Experiment was conducted in the climate- controlled greenhouse with an automated sprinkler system in November, 2015. A day/night temperature of 78oF/75oF was maintained in greenhouse with 15% shade. The overhead sprinkler system was set up for 1 min/12 hours. Healthy Viburnum (Viburnum obovatum – Mrs. Schiller’s delight variety) rooted cuttings grown in potting mix #2 (Morton’s Horticultural Products, Inc. McMinnville, TN, USA) was used for the experiment. No additional fertilizer was applied during experiment. The pot size was 10 cm X 10 cm and pots were inoculated with the tree inoculum levels of the three inoculation methods. Three single-plant replications per inoculum level were arranged in a randomized complete block design. Non-inoculated sterilized soil served as a control.

Data collection

Plants were evaluated for disease severity 28 days after inoculation using a 1-5 ordinal scale : 1 = no symptom, heavily branched root system, and healthy looking; 2 = light brown necrosis in distinct spots, often necrosis in the root tip, less branched root system than healthy roots; 3 = few side roots, and dark brown necrosis in distinct spots; 4 = few and small side-roots, and dark brown necrosis of most of the root system, or all around the stem; 5 = plant dead. Plant and root fresh weight was recorded at the end of the experiment. Disease severity scales were converted into percentage (0-100% of roots affected) before analyzing.

Statistical analysis: Inoculation method and inoculum level effect on disease severity was determined using the general linear model procedure of SAS (PROC GLM, SAS version 8.2, SAS Institute, Carry, NC) and means were separated using Fisher’s least significant difference test.

Objective 1.2. Pathogenicity of Phytophthora nicotianae and Rhizoctonia solani to biofumigant cover crops.

Germination percentages of Brassicaceae cover crops

Ten seeds from the each tested cover crop were placed on each petri plate with #1 filter paper on 3 May 2016. Prior placing the seeds on the filter paper, filter papers were wetted with 5 ml of steilized water to facilitate germination. Germination count was taken on 5, 9 and 11 May, 2016 and counts were converted into percentages using the following formula before the data analysis. Germination percentage= No of seeds germinated / Total number of seeds planted *100% (Pirasteh-Anosheh and Hamidi 2013).

Fungal cultures and inoculum preparation

Phytophthora nicotianae and Rhizoctonia solani cultures were obtained from the culture collection of Dr. Fulya Baysal-Gurel at Tennessee State University Otis L. Floyd Nursery Research Center McMinnville, TN. Isolate FBG201505 of R. solani, originally isolated from a diseased Viburnum plant, and isolate FBG201506 of P. nicotianae, originally isolated from a diseased Hydrangea plant, cultures were maintained on potato dextrose agar and V8 agar medium, respectively. Prior to the study R. solani was inoculated into Viburnum plant roots and P. nicotianae was inoculated Hydrangea roots and subsequently recovered from the diseased roots to ensure virulence of the pathogens. Rice grain inoculum (Holmes and Benson 1994) was prepared by autoclaving 25 g long grain rice and 18 ml deionized water in a 250 ml flask twice consecutively and three 7-mm V8 agar plugs colonized by P. nicotianae were placed in the flask. Infected rice grains were grown 2 weeks and mixed weekly prior to use as inoculum. Seven days old R. solani culture was decanted into a sterile beaker with sterile distilled H2O (1,000 ml) and homogenized with blender (Hamilton Beach hand blender, Model number 59785R) to make slurry at 1 petri dish/L inoculum level.

Pathogenicity testing of Rhizoctonia solani and Phytophthora nicotianae on Brassica cover crops.

Brassica seeds from 15 different Brassica cover crops (Table 2.2) were sown into 10-cm pots containing autoclaved topsoil (soil provider- McMinnville Lawn & Garden LLC, TN, USA). The selection of Brassica cover crops for the experiments was based on a number of factors such as their commercial availability as biofumigant crops or their high level of effective ITC-producing GSLs (Lord et al. 2011).

Ten seeds were added into each pot and pots were inoculated with previously described inoculation methods. R. solani experiment trials were started on March 16, 2017 and P. nicotianae trials were started on May 1, 2017. Experiments repeated twice for P. nicotianae and R. solani. One P. nicotianae infested rice grain was placed in each pot based on previous study and gently mixed to incorporate with the soil (Benson et al. 1997). Control pots received a rice grain prepared with V8 agar plugs with no pathogen presence. Fifty milliliters of the R. solani agar slurry, prepared as described earlier, was used to inoculate each 10-cm pot. Control pots received 50 ml of agar slurry prepared with PDA plates with no pathogen presence. Each pot containing 10 cover crop seeds was considered as one experimental unit, with 6 pots per each cultivar per each pathogen. All the plants in each experiment were inoculated in the same day and plants were watered using sprinkler system 1 min/12 hours to maintain adequate soil moisture. Pre- emergence and post emergence damping off and disease symptoms were recorded every week. Plant height from the soil line to the tallest expanded foliage, plant widths as perpendicular lines, plant fresh weight and root weight were measured at the end of the trial. Roots were rinsed in tap water to remove adhering soil and approximately 50% of the plants of each cover crop were selected for the pathogen re-isolation. Necrotic or water soaked tissues were rinsed in sterile water and was surface sterilized in 70% ethanol for 5 sec prior planting on V8 PARPH (Appendix 1.5) or R. solani selective media (Appendix 1.2) and pathogens were re-isolated form the cultures. The experiments were arranged as complete randomized designs and were conducted twice.

Statistical analysis

Plant growth data, root disease severity and percentages was determined using the general linear model procedure of SAS (PROC GLM, SAS version 8.2, SAS Institute, Carry, NC) and means were separated using Fisher’s least significant difference test.

Table 2. Brassica cover crops evaluated in the pathogenicity experiment

Crop

Scientific name

Seed rate

Price/lb

Days to maturity

Intended use

Oilseed radish

Raphanus sativus

25 lb/A

$8.35

60-90

Biofumigation

Mustard

Sinapis alba

15-20 lb/A

$4.95

80-90

Biofumigation

Purple top forage turnips

Brassica rapa

5-8 lb/A

$7.45

60-90

Fresh market

Arugula

Eruca vesicaria spp. sativa

30-50 seeds/ft2

$23.80

21 baby, 40 full

Fresh market

Sylvetta green arugula

Diplotaxis tenuifolia

30-50 seeds/ft2

$107.95

50

Fresh market

Wasabi arugula

D.erucoides

30-50 seeds/ft2

$212.20

45

Fresh market

Bellezia arugula

D. erucoides

30-50 seeds/ft2

$189.25

51

Fresh market

Dragon’s tongue arugula

D. tenuifolia

30-50 seeds/ft2

$216.40

50

Fresh market

Olive-leaved sylvetta arugula

D.s tenuifolia

30-50 seeds/ft2

$99.55

50

Fresh market

Astro arugula

E. vesicaria spp. sativa

30-50 seeds/ft2

$29.45

38

Fresh market

Mighty mustard® pacific gold

B. juncea

15-20 lb/A

$8.60

80-90

Biofumigation

Oriental mustard

B.juncea

15 seeds/ft2

$3.67

4-5

Condiment industry

Red giant mustard green

B. juncea

15 seeds/ft2

$52.95

21 baby 45 full

Fresh market

Dwarf essex rape

B. napus

5-10 lb/acre

$6.15

60-80

Biofumigation

Amara mustard green

B. carinata

15 seeds/ft2

$32.90

21 baby 40 full

Fresh market

Objective 2. Determining the effect of biofumigation in suppressing Rhizoctonia solani and Phytophthora nicotianae in woody ornamental nurseries

Brassicaceae cover crops and inoculum preparation. The experiments were carried out in the greenhouse at the Otis L. Floyd Nursery Research Center in McMinnville, TN.

Eight Brassicaceae cover crops were planted to study their potential biofumigant activity and possible suppressive effect on Rhizoctonia solani and Phytophthora nicotianae (Table 1). Crops were selected based on findings of previous pathogenicity experiments (Liyanapathiranage 2017). The seeding rates were based on the seed suppliers’ recommendation per acre. Pots used were 16 cm in height, 16 cm in diameter, and filled with 2.5 kg of sterilized sandy loam topsoil using electric soil sterilizer (Pro-Grow Supply Corp., Model SS-30 Brookfield, WI, USA). There were six replicates of each treatment for each pathogen, P. nicotianaeor R. solani. Two types of controls were maintained, inoculated without cover crops and non-inoculated without cover crops. Experiments were arranged in a completely randomized design (CRD) with six replicates and conducted twice, 20 June and 27 October in 2016.

Isolate FBG201506 of Phytophthora nicotianae andisolate FBG201505 of Rhizoctonia solaniwere obtained fromthe culture collection of Dr. Fulya Baysal-Gurel at the Tennessee State University Otis L. Floyd Nursery Research Center McMinnville, TN. The R. solanispecimen was originally isolated from a diseased viburnum (Viburnum odoratissimum) plant and maintained on PDA medium. The P. nicotianae specimen was originally isolated from a diseased hydrangea (Hydrangea paniculata)plant and maintained on V8 agar medium. Prior to this study, viburnum plant roots were inoculated with the R. solani culture while hydrangea plant roots were inoculated with the P. nicotianae culture and subsequently re-isolated from the diseased roots to confirm pathogen virulence.

For P. nicotianaeinoculum, rice grain inoculum technique, modified after Holmes and Benson (1994), was followed. Briefly, 25 g of long grain rice in 18 ml deionized water was autoclaved twice, three 7-mm sized P. nicotianaecolonized V8 juice agar (100 ml of clarified V8 juice, 15 g of agar, and 900 ml of deionized H2O) plugs were placed in the 250-ml flask and incubated for two weeks at room temperature. The inoculum in the flask was mixed weekly prior to a final use. Four P. nicotianaecolonized rice grains were added onto each pot for inoculation. For R. solaniinoculum, agar slurry was prepared (1 plate of a 7-day-old R. solaniculture blended with 1L of sterilized distilled water) and each pot drenched with 100 ml slurry; this volume of inoculum was determined by conducting a preliminary experiment with dried sterilized topsoil. All pots in each experiment were inoculated and seeded on the same day. Plants were watered using a sprinkler system for 1 minute every 12-hrs to maintain adequate soil moisture. All the flowering cover crops were up-rooted on 12 December 2016 for the first experiment and on 20 February 2017 for the second experiment.

Preparation of green manure and incorporation. Plant stems, leaves, flowers and roots of the cover crops were harvested, chopped into approximately 1 cm2 pieces, and immediately incorporated into the soil.  One hundred and twenty five grams of plant material including fresh shoots and roots was incorporated into each pot to a depth of 15 cm. After incorporation, pots were covered with a sheet of polyethylene (clear polyethylene sheeting, 2.5 m x 30 m x 3 mm thickness, Wrap Bros, Chicago, IL, USA) for 2 weeks and 4 weeks to test the effect of biofumigation time on cover crop phytotoxicity to the fallow crop (Goud et al. 2013). Soil temperature and moisture in the pots were measured using Watchdog meters (Spectrum technologies, Model 150, Aurora, IL, USA). Soil samples were taken prior to incorporation of cover crops with a biofumigant property and volatiles were collected on a daily basis to evaluate the effect of biofumigation on the soil pathogen community.

Volatile collection and analysis. During biofumigation, volatiles were collected from the covered pots at varying time intervals (24-hrs, 1 week, 2 weeks, 4 weeks). Volatiles were collected using a 30 cm long, 0.6 cm diameter hollow metal probe inserted into the soil half way to the bottom of the pot. Volatiles from the soil were collected on adsorbent filters filled with 50 mg of Super-Q glass traps (lltech Associates Inc, Illinois, USA) attached between the metal probe and LaMotte® Air Sampling vacuum pump (LaMotte, MD, USA) using inert Tygon® tubing. Volatiles from the soil airspace were collected with an air suction flow of 0.25 LPM. Volatile collections were performed on each pot for 2 hrs. The Super-Q traps were then eluted with 200 µl of methylene chloride using active pressure from the pipette and purified nitrogen gas into 2 ml clear glass vials (Agilent Technologies, Part Number 5182-0715 Santa Clara, CA, USA) equipped with 250 µl glass inserts (Agilent Technologies, Part Number 8010-0132 Santa Clara, CA, USA) (Ali et al. 2012). Glass vials with extracts weresealed with Teflon tape and stored in a freezer at -20°C until analyzed on a gas chromatograph mass spectrometer (GC-MS).

The volatile analysis was carried out with a Shimadzu QP 2010 GC-MS (Shimadzu Corporation, Japan) equipped with a DB-5ms capillary column (30 m length x 0.25 µm film thickness x 0.25 mm diameter) (Agilent Technologies Santa Clara, CA, USA). Helium was used as the carrier gas at 7.8 psi, with a flow rate of 1 ml/min with a 3 ml/min purge flow. The injector temperature was 220°C, the injection volume was 1 µl, and all injections were splitless. The GC ovenwas held at 50°C for 2 min and raised 10°C/min to 260°C and held for 5 min. The mass spectra were acquired with a source temperature of 230°C under a -70eV ionization potential.  Full scan analysis was performed in the mass range 45-500m/z. Results were evaluated by LabSolutions software (Shimadzu Corporation, Japan).Inoculated, non-fumigated pots were used as controls in order to exclude non-biofumigant related volatiles. Remaining compounds were tentatively identified by comparing retention times with the mass spectral database (NIST08).

Woody ornamental crop bioassay. Viburnum (Viburnum odoratissimum) and hydrangea (Hydrangea paniculata) rooted cuttings were planted in the treatment pots  4 weeks after biofumigation in the first experiment, and 2 weeks after biofumigation in the second experiment. Viburnum  rooted cuttings (Liner source Inc, Eustis, FL) were transplanted into the pots that had been infested with Rhizoctonia solanion 24 January 2017 and 3 July 2017, and hydrangea  rooted cuttings (Schaefer Nursery Co., Winchester, TN) were transplanted into the pots that had been infested with Phytophthora nicotianae on 25 January 2017 and 7 March 2017. Plant height and widths (maximum width; the widest part from leaf tip to leaf tip) were measured immediately after transplantation and experiment termination. Plant total fresh weight and root fresh weight were also determined for each viburnum and hydrangea experimental plants immediately at the end of the experiments. Plants were monitored for phytotoxicity symptoms throughout the experiments

The severity of root rot was assessed using a scale of 0−100% roots affected at the end of the experiments. Forty root pieces (1-cm sized) were cut from each root ball and were cultured on selective media, ten pieces per media plate with 4 replications. Viburnum roots were cultured on Rhizoctonia selective medium (Gutierrez et al. 2001) while hydrangea root pieces were cultured on PARPH-V8 medium (Ferguson and Jeffers 1999). Plates were incubated at 25°C in the dark conditions (VWR incubator, PA, USA).  The number of root pieces showing Rhizoctonia growth was counted after 48-hrs, while Phytophthora growth was counted after 5 days.

Determination and quantification of microorganisms in the soil. Soil samples were taken from each pot at the beginning and end of the experiments and they were stored at 4°C until they could be cultured. Samples were processed and serial diluted. Stock solution and 10-1serial dilution was cultured on Rhizoctonia or Phytophthora selective or semi-selective media (Gould et al. 1985). Rhizoctonia plates were incubated at 25°C for 48 to 72-hrs under dark conditions. Phytophthoraplates were incubated at 25°C for 5 days under dark conditions. Colony counts were expressed as colony forming units (CFU) per gram of dried soil. Randomly selected colonies were re-cultured on respective media and identified using DNA sequencing. DNA extraction was performed using a PowerLyzer®UltraClean® microbial DNA isolation kit with the MOBIO laboratories Inc. protocol. DNA quantification was done with NanoDrop 2000c (Thermo Scientific, Wilmington, DE, USA). PCR was conducted using ITS1 and ITS4 primers. Positive PCR products identified from the agarose gel electrophoresis, were purified using Wizard® SV gel and PCR clean up system procedure (Promega corporation, Madison, WI, USA) and sent for DNA sequencing to Eurofins genomics (Eurofins genomics, KY, USA).

Soil tests. Soil samples taken from each pot including control pots, at the end of the experiment were analyzed for pH, phosphorus, potassium, calcium, magnesium, organic matter and nitrogen content by sending the samples to the University of Tennessee Extension Institute of Agriculture – Soil, Plant and Pest Center in Nashville, TN, USA.

Data analysis. Plant height and width, fresh weight, root weight, root rot disease severity and colony counts data were analyzed using the general linear model (PROC GLM) procedure of SAS (version 8.2, SAS Institute, Carry, NC). Means were separated using Fisher’s least significant difference test and correlations were determined by Spearman linear regression analysis.

Research results and discussion:

Objective 1. Evaluating pathogenicity of Rhizoctonia solani and Phytophthora nicotianae to Brassicaceae biofumigant cover crops

Evaluation of inoculation methods and inoculum level of Rhizoctonia solani to determine disease response.

Rhizoctonia solani-infested viburnum plants displayed dark-colored, rotten primary and secondary roots. Compared with healthy roots, inoculated root growth was restricted and overall volume was low; outer cortex of roots was invaded by infectious hyphae.

Inoculation methods of R. solani had a significant negative correlation with plant fresh weight (R2=-0.509, P=0.001) (Data not shown). Lowest fresh weight was recorded for the viburnum plants inoculated with the agar slurry method at 2 petri dishes/L inoculum level (Table 3). All the tested inoculation methods at three different inoculum levels had significantly higher root rot disease percentages compared to non-inoculated control (P = 0.00). Viburnum plants inoculated with the agar slurry method at 3 petri dishes/L inoculum level showed the most severe root rot disease, but the difference between the three levels was not statistically significant. All three inoculum levels of agar slurry method had significantly higher root rot disease severity compared to the other tested inoculation methods as well as the non-inoculated control. Although agar plugs and chopped potato soil medium inoculated plants had significantly greater root rot disease percentages compared with non-inoculated control plants, their plant fresh weights were similar to the non-inoculated control plants (Table 3).

Pathogenicity of Phytophthora nicotianae and Rhizoctonia solani to Brassicaceae covercrops.  

In in vitro experiment, 100% seed germination was observed for the olive-leaved sylvetta green, oilseed radish, and astro arugula two days after seeding. Lowest seed germination after 2 days was recorded by amara mustard green (73.3%), red giant mustard green (66.7%) and dragon’s tongue arugula (50.0%). When observations were taken 6 days after seeding oriental mustard, dwarf essex rape and purple top forage turnips also recorded 100% germination while bellezia arugula (80.0%), red giant mustard green (76.6%) and dragon’s tongue arugula (73.3%) had lowest seed germination. Eight days after seeding, none of the cover crops showed increased germination from 6 days (data not shown). Since there were differences on seed germination percentages between the cover crops in in vitro experiment, non-inoculated control plants were included in the pathogenicity test to determine accurate disease incidence percentages.

Both of the tested soilborne pathogens had similar disease symptoms such as pre- and post-damping off, stunting, chlorosis, wilting and plant death. The disease symptoms and signs of the pathogens were observed on the root system and root collar areas one month after inoculation. Brassicaceae cover crops that were grown on R. solani infested soil showed disease severity percentages ranging from 7.7% to 85.9% and disease incidence percentages from 0.7% to 100.0% (Table 4). Cover crops that were inoculated with P. nicotianae had disease severity percentages ranging from 1.1% to 77.9% and disease incidence percentages between 0.0% to 100.0%. Among the Brassicaceae cover crops tested with R. solani; arugula, wasabi arugula, bellezia arugula, dragon’s tongue arugula and olive-leaved sylvetta arugula showed significantly higher root rot severity compared to the other tested cover crops (P = 0.00) and their disease severity percentages were above 77.6%. Among the Brassicaceae cover crops tested with P. nicotianae; bellezia arugula and olive-leaved sylvetta arugula showed significantly higher root rot severity compared to the other tested cover crops (P = 0.00) and their disease severity percentages were above 65.1%. 

A negative correlation was observed between plant height, width and fresh weight with root rot disease incidence and severity caused by both pathogens. (Table 5). Brassicaceae cover crops inoculated with R. solani had significant height reduction percentages (P = 0.003) and fresh weight reduction percentages (P = 0.037) while plant width was not affected (P = 0.070). Height (P = 0.248), width (P = 0.702) and weight (P = 0.059) reduction percentages were not significant in the plants that were inoculated with P. nicotianae. The greatest height reduction percentage was recorded by sylvetta green plants that were inoculated with R. solani (30.0% reduction) and P. nicotianae (27.3% reduction). Dragon’s tongue arugula plants had the greatest fresh weight reduction (32.0%) when inoculated with P. nicotianae, and bellezia arugula plants had the greatest fresh weight reduction (44.5%) when inoculated with R. solani. Purple top forage turnips had the least fresh weight reduction percentage under R. solani disease pressure, while oilseed radish had the least fresh weight reduction under P. nicotianae disease pressure (Table 6). Among the tested Brassicaceae crops that are commercially used for biofumigation (Table 2), mustard (Sinapis alba) showed significantly lower fresh weight when inoculated with R. solani compared to control plants and P. nicotianae (Figure 1). 

Root rot disease severities were significantly higher in oilseed radish, purple top forage turnips, wasabi arugula, dragon’s tongue arugula and arugula (Eruca vesicaria subsp. sativa) Brassicaceae crops inoculated with R. solani compared to the plants inoculated with P. nicotianae (Figure 2).

Discussion

Rhizoctonia solani is a very common soilborne fungus capable of infecting great diversity of plants from early germination and seedling stage (pre- and post-emergence damping-off) through mature plant stage (root and crown rots) (Marasus and Bredell 1973). Most of the woody ornamental species are particularly susceptible to infection by Rhizoctonia, Phytopththora and Pythium species during the initial production stages (Goss 1978). 

These soilborne pathogens can kill cuttings and cause major economic losses to woody ornamental nurseries. Variability in disease symptom expression makes it difficult to identify the diseases, and when diseases are unidentified or incorrectly identified, it can result in passing of infected plants to the other stages of the nursery production (McCully and Thomas 1977). Identifying an effective inoculation method for a pathogen is crucial in order to test control practices. Attempts to identify resistant varieties to R. solani are difficult due to the lack of artificial disease exposure techniques that produce highly uniform disease distributions (Pierson 1959). In this study, we were able to identify an agar slurry method as an effective inoculation method to infect woody ornamental viburnum plants with R. solani. Since there were no significant differences between all three inoculum levels of the agar slurry method in Rhizoctonia root rot disease severity, when conducting future experiments using agar slurry method, the lowest inoculum level of 1 petri dish/L can be used to get a sufficient disease response and will minimize resource use compared to 2 or 3 petri dishes/L inoculum level as we used in pathogenicity tests. This method has subsequently been used to evaluate biological products and fungicides to control R. solani on viburnum plants under field conditions and has demonstrated disease severity percentage up to 63.0 and 66.4% in inoculated, non-treated plots in 2016 and 2017, respectively (Baysal-Gurel et al. 2017 and 2018). 

Cover crops in general provide a wide range of benefits in production systems: they suppress weeds, reduce erosion, preserve soil quality, and provide organic matter and aid in nutrient cycling (Mutch and Snapp 2003). Plants belonging to the Brassicaceae family are widely cultivated as important vegetable crops, for oilseed, as condiments and forage crops (Fenwick et al. 1983). Along with other families in the order Capparales, Brassicaceae are comprised of plants containing glucosinolate (GSL) compounds (Fenwick et al. 1983; Brown and Morra 1997). Glocosinolates are secondary plant products containing sulfur compounds, which can control soilborne pathogens upon hydrolysis into isothiocyanates (ITC).

Matthiessen and Kirkegaard (2006) and Motisi et al. (2010) have compiled a list of pathogens that are suppressed by biofumigant plants. Some of the important groups of pathogens that have been targeted including Phytophthora, Aphanomyces, Pythium, Fusarium, Gaumannomyces, Sclerotinia, Rhizoctonia and Verticillium. Brassica plant species and cultivars have different types, concentrations and distributions of GSL compounds in their plant parts (Josefsson 1967; Sang et al. 1984) and the ability to produce ITC may vary from one species to another. Sensitivity of soilborne fungal and oomycete pathogens to these different types and concentrations of ITC vary (Brown and Morra 1997). These variations in sensitivity to ITC generate opportunities to select Brassica species among its 2,500 species which have the largest quantities of ITC that are most toxic to the target pathogens. 

Cover crops are usually direct-seeded into the field, grown until flowering and then incorporated into the soil prior to the planting of an economically important crop (Snapp et al. 2006). Therefore, the ability of the cover crop to germinate and survive in a field that is heavily infested by soilborne pathogens is crucial to successful biofumigation. Infected cover crops will suffer pre- and post-damping-off, wilting, chlorosis, stunting and plant death (Krasnow and Hausbeck 2015). Delayed seedling emergence increases the opportunity for soilborne disease infection and the probability of pre-emergence damping-off (Peace 1962) and further delays in attack by pathogens, results post-emergence damping-off. While the seedling is still in the juvenile stage, symptoms may develop through the soil surface and the susceptibility of the seedling may be reduced with seedling maturity and lignification. Increased resistance may be due to pectin conversion to calcium pectate in plant tissues, which increases tissue resistance to the polygalacturonase enzyme released by pathogens (Bateman and Lumsden 1965).

In this experiment, 15 varieties of Brassicaceae cover crops were tested against R. solani and P. nicotianae. All the tested arugula varieties were intended for fresh market, and except for astro arugula, all species had higher root rot disease severity when grown in soil inoculated with both soilborne pathogens. Oilseed radish, mustard, mighty mustard® pacific gold and dwarf essex rape are commercially used for biofumigation, and in this study displayed relatively low root rot disease percentages when inoculated with either R. solani or P. nicotianae. These five Brassicaceae cover crops also had the highest fresh weights compared to the other tested Brassicaceae crops, which is an important factor for the biofumigation efficiency. 

Although purple top forage turnips, astro arugula, oriental mustard and amara mustard green are not currently utilized as biofumigation cover crops, they also exhibited lower root rot disease severity and higher fresh weights.

Antonious and his colleagues identified Brassica napus (winter rapeseed) as a crop with higher above ground biomass (7.7 ± 3.4 kg m2) (Antonious et al. 2009). Despite a relatively low GSL content (27 ± 14 µmoles g-1), its higher biomass resulted in a higher GSL yield per unit area. Raphanus sativus (oilseed radish) can produce approximately 10 kg m2 of biomass, and also contains higher quantities of GSL (Sundermeier 2008). In this study significantly higher root rot disease severities and higher plant fresh weights were observed in oilseed radish, when inoculated with R. solani. 

R. solani is known to cause both pre-emergence and post-emergence damping-off in crucifers and other crops (Baird 1996). Previous studies have shown R. solani infections can causes seedling diseases in oilseed rape at a rate of 80-100% and yield loss up to 30% (Tahvonen et al. 1984; Khangura et al. 1999). 

After using Brassica seed for oil production, defatted seed meal is produced as a byproduct (e.g. mustard meal). This seed meal contains high amounts of GSL compounds and the myrosinase enzyme required for hydrolysis to ITC, so it could be used as a biofumigant soil amendment (Brown and Morra 1997).  Previous studies have shown these materials effective against major soilborne pathogens, including Rhizoctonia species (Mazzola et al. 2007). Liquid formulations and dried pellets developed from high GSL plants have also shown inhibition activity in vitro against Meloidogyne incognita (De Nicola et al. 2013), as well as Rhizoctonia and Pythium (Lazzeri et al. 2004).

Dried brown mustard plants are reported to be effective at preserving GSLs and myrosinase and as reported by Michel (2014) and also were able to significantly reduced Verticillium dahlia microsclerotia in greenhouse soil. This method of biofumigant incorporation is important during periods of the year like winter, when biofumigant plant growth is restricted and will be highly important when the cover crop itself is highly susceptible to the pathogens existing in the field.

As a result of this study, oilseed radish, mustard, purple top forage turnip, astro arugula, mighty mustard® pacific gold, oriental mustard, dwarf essex rape Brassica, amara mustard green had low root rot disease severity and incidence in soil which had pre-existing populations of R. solani or P. nicotinanae under greenhouse conditions. Currently, astro arugula, oriental mustard and amara mustard green are produced solely for the fresh market, but since these Brassicaceae crops can withstand a higher disease pressure from R. solani and P. nicotinanae, further investigation is warranted alongside other cover crops already being used for biofumigation.

LITERATURE CITED

Anosheh, H. P., Sadeghi, H., and Emam, Y. 2011. Chemical priming with urea and KNO3 enhances maize hybrids (Zea mays L.) seed viability under abiotic stress. Journal of Crop Science and Biotechnology, 14(4), 289-295.

Antonious, G. F., Bomford, M., and Vincelli, P. 2009. Screening Brassica species for glucosinolate content. Journal of Environmental Science and Health Part B 44: 3, 311-316.

Baird, R. E. 1996. First report of Rhizoctonia solani AG-4 on canola in Georgia. Plant Dis. 80:10.

Baker, K. F. 1970. Types of Rhizoctonia diseases and their occurance, p. 125-148. In: J. R. Parmeter, Jr. (eds.). Rhizoctonia solani, biology and pathology. University of California Press, Los Angeles, CA.

Bateman, D. F., and Lumsden, R. D. 1965. Relation of calcium content and nature of the pectic substances in bean hypocotyls of different ages to susceptibility to an isolate of Rhizoctonia solani. Phytopathology 55:734-738.

Baysal-Gurel, F., Simmons, T., Liyanapathiranage, P., Kabir, Md. N. 2017. Evaluation of biorational products and fungicides for the control of Rhizoctonia root rot of viburnum, 2016. Plant Disease Management Report No. 11: OT003. 

Baysal-Gurel, F., Simmons, T., Kabir, Md. N. 2018. Evaluation of biological products and fungicides for the control of Rhizoctonia root rot of viburnum, 2017. Plant Disease Management Report No. 12: OT025. 

Benson, D. M. and Cartwright, D. K. 1996. Ornamental diseases incited by Rhizoctonia spp., p. 303-314. In: B. Sneh, S. Jabaji-Hare, S. Neate, and G. Dijst (eds.). Rhizoctonia species: Taxonomy, molecular biology, ecology, pathology and disease Control. Kluwer, Dordrecht, The Netherlands.

Benson, D. M., Hinesley, L. E., Frampton, J., and Parker, K. C. 1998. Evaluation of six Abies spp. to Phytophthora root rot caused by Phytophthora cinnamomi. Biol. Cult. Tests Control of Plant Dis, 13, 57.

Berkenkamp, B., Vaartnou, H. 1972. Fungi associated with rape root rot in Alberta. Can. J. Plant Sci. 52:973–976.

Blair, J. E., Coffey, M. D., Park, S. Y., Geiser, D. M., Kang, S. 2008. A multi-locus phylogeny for Phytophthora utilizing markers derived from complete genome sequences. Fungal Genetics and Biology 45: 266-277.

Blum, L. E. B. and Rodríguez-Kábana, R. 2006. Dried powders of velvet bean and pine bark added to soil reduce Rhizoctonia solani– induced disease on soybean. Fitopatologia Brasileira 31:261-269. 

Brown, P. D., Morra, M. J., 1997. Control of soilborne plant pests using glucosinolate-containing plants. In: Donald LS, ed. Advances in Agronomy Volume 61. Academic Press, 167-231.

Cacciola, S. O., Magnano di San Lio, G. 2008. Management of citrus diseases caused by
Phytophthora spp. In: Ciancio, A., Mukerji, K. G. (eds), Integrated management of diseases caused by fungi, phytoplasma and bacteria. Springer Science+Business Media, pp. 61-84.

Carling, D. E and Sumner, D. R. 1992. Rhizoctonia. In methods for research on soil borne phytopathogenic fungi (eds. Singleton, L. L., Mihail, J. D. & Rush, C. M.).
American Phytopathological Society. 157–165.

Daughtrey, M. L., and Benson, D. M. 2005. Principles of plant health management for ornamental plants. Annual Review of Phytopathology 43:141-169.

De Nicola, G. R., D’avino, L., Curto, G., Malaguti, L., Ugolini, L., Cinti, S., Patalano, G., Lazzeri, L., 2013. A new biobased liquid formulation with biofumigant and fertilizing properties for drip irrigation distribution. Industrial Crops and Products 42, 113-8.

Dolmans, N., and Looman, B. H. M. 1984. Rhizoctonia (dradenschimmel) – bestrijding. (With English summary.) Pages 169-173 in: Jaarboek 1984. Proefstation voor de Boomkwekerij, Boskoop, The Netherlands.

Dreistadt, S. H. 2001. Integrated pest management for floriculture and nurseries. Publication #3405, ANR Publications, University of California, 6701 San Pablo Avenue, Oakland, CA 94608-1239. 422 pp.

Ellis, P. J. 1983. Diseases of rapeseed in the Peace River region of Alberta. M.P.M. Thesis. Simon Fraser University, Burnaby, B.C., Canada: 89 p.

Erwin, D. C., Ribeiro, O. K., 1996. Phytophthora diseases worldwide. St. Paul, MN, APS Press. Ferguson, L., Sakovich, N., and Roose, M. 1990. California citrus rootstocks. University of California Division of Agriculture and Natural Resources, Oakland. Publication 21477.

Farr, D. F., Bills, G. F., Chamuris, G. P., and Rossman, A. Y. 1989. Fungi on plant and plant Products in the United States. APS Press, St. Paul, MN. p. 496.

Fenwick, G. R., Heaney, R. K., and Mullin, W. J. 1983. Glucosinolates and their breakdown products in food and food plants. Crit. Rev. Food Sci. Nutr. 18, 123-201.

Ferguson, A. J., and Jeffers, S. N. 1999. Detecting multiple species of Phytophthora in container mixes from ornamental crop nurseries. Plant Dis. 83:1129. 

   Garibaldi, A., & Gullino, M. L. (1990). Disease management of ornamental plants: a never ending challenge. Disease management of ornamental plants: a never-ending challenge., 55(2a), 189-201.

Goss, O. M. 1978. Pathogens in plant propagation. comb.  Proceeding. Int. Plant Propagators Soc. 28:400-406.

Gupta, D. K. 1985. Root and collar rot of mustard in Sikkim. Indian J. Mycol. Plant Pathol. 15: 225.

Gutierrez, W. A., Shew, H. D., Melton, T. A. 2001. A semi-selective medium to isolate Rhizoctonia solani from soil and tissue. Plant Pathological Extension, 1-2.

Hall, B., Barlow, T., Rawnsley, B., Hitch, C., and Deland, L. 2008. Varietal resistance of cauliflower cultivars to soil borne diseases: Rhizoctonia solani and Leptosphaeria
maculans. www.pir.sa.gov.au

Holmes, K. A., and Benson, D. M. 1994. Evaluation of Phytophthora parasitica var. nicotianae for biocontrol of Phytophthora parasitica on Catharanthus roseus. Plant disease, 78(2), 193-199.

Houston, B. R. 1945. Culture types and pathogenicity of isolates of Corticium solani.
Phytopathology. 35: 371–393.

Hu, J. H., Hong, C. X., Stromberg, E. L., and Moorman, G. W. 2008. Mefenoxam sensitivity and fitness analysis of Phytophthora nicotianae isolates from nurseries in Virginia, USA. Plant Pathol. 57:728-736. https://doi.org/10.1111/j.1365-3059.2008.01831.

Huber, D. M., Christmas, E. P., Herr, L. J., McCay-Buis, T. S., Baird, R. 1992. Rhizoctonia crown rot of canola in Indiana. Plant Dis. 76: 1251–1253.

Jiskani, M. M., Pathan, M. A., Wagan, K. H., Imran, M., and Abro, H. 2007. Studies on the control of tomato damping-off disease caused by Rhizoctonia solani Kuhn.
Pakistan Journal of Botany. 39(7): 2749-2754.

Johnson, E. S., Wolff, M. F., Wernsman, E. A., Rufty, R. C., 2002. Marker-assisted selection for resistance to black shank disease in tobacco. Plant Disease 86: 1303-1309.

Jones, R. K., and Benson, D. M., eds. 2001. Diseases of woody ornamentals and trees in nurseries. American Phytopathological Society, St. Paul, MN.

Josefsson, E. 1967. Distribution of thioglucosides in different parts of Brassica plants. Phytochemistry, 6, 1617-1627.

Judelson, H. S., Blanco, F. A., 2005. The spores of Phytophthora: weapons of the plant destroyer. Nature Reviews Microbiology 3, 47–58.

Kamoun, S. 2006. A catalogue of the effector secretome of plant pathogenic oomycetes. Annual Review of Phytopathology 44:41–60.

Khangura, R. K., Barbetti, M. J., Sweetingham, M.W. 1999. Characterization and pathogenicity of Rhizoctonia species on canola. Plant Dis. 83 714–721. 10.1094/PDIS.1999.83.8.714.

Ko, W. H., and Hora, F. K. 1971. A selective medium for the quantitative determination of Rhizoctonia solani in soil. Phytopathology, 61(6), 707-710.

Krasnow, C. S., and Hausbeck, M. K. 2015. Pathogenicity of Phytophthora capsici to Brassica vegetable crops and biofumigation cover crops (Brassica spp.). Plant Disease, 99(12), 1721-1726. DOI: 10.1094/PDIS-03-15-0271-RE.

Lazzeri, L., Leoni, O., Manici, L. M. 2004. Biocidal plant dried pellets for biofumigation. Industrial Crops and Products 20, 59- 65.

Lichtenzveig, J., Anderson, J., Thomas, G., Oliver, R and Singh, K. 2006. Inoculation and growth with soil borne pathogenic fungi. Medicago truncatula handbook. 1-10.

Lord, J. S., Lazzeri, L., Atkinson, H. J., and Urwin, P. E. 2011. Biofumigation for control of pale potato cyst nematodes: activity of brassica leaf extracts and green manures on Globodera pallida in vitro and in soil. Journal of agricultural and food chemistry, 59(14), 7882-7890.

MacNish, G. C., Carling, D. E., Sweetingham, M. W., Ogoshi, A and Brainard, K. A. 1995. Characterization of anastomosis Group-10 (AG-10) of Rhizoctonia solani.
Australasian Plant Pathology. 24: 252–260.

Marasus, W. F. O., and Bredell, I. H. 1973. Microflora of South African lucerne (Medicago sativa L.) seed. Phytophylactica 5: 89-94.

Matthiessen, J. N., and Kirkegaard, J. A. 2006. Biofumigation and enhanced biodegradation: opportunity and challenge in soilborne pest and disease management. Critical Reviews in Plant Sciences, 25: 235-265.

Mazzola, M., Brown, J., Izzo, A. D., Cohen, M. F. 2007. Mechanism of action and efficacy of seed meal-induced pathogen suppression differ in a Brassicaceae species and time-dependent manner. Phytopathology 97, 454-460.

McCully, A. J., and Thomas, M. B. 1977. Soilborne diseases and their role in plant propagation. Comb. Proc. Int. Plant propagators Soc.27:339-350.

Menzies, J. D. 1970. Introduction: The first century of Rhizoctonia solani, p. 3-5. In: J. R. Pa rmeter, Jr. (eds.). Rhizoctonia solani, biology and pathology. University of California Press, Los Angeles, CA.

Michel, V. V. 2014. Ten years of biofumigation research in Switzerland. Aspects of Applied Biology 126, 33-42.

Mizell, R., Knox, G., Knight, P., Gilliam, C., Arthers, S., Austin, R., Baldwin, H. 2012. Woody ornamental and landscape plant production and pest management innovation strategic plan. USDA Southern Region IPM Center, North Carolina State University, Raleigh.

Moralejo, E., Pérez-Sierra, A. M., Álvarez, L. A., Belbahri, L., Lefort, F., Descals, E. 2009. Multiple alien Phytophthora taxa discovered on diseased ornamental plants in Spain. Plant Pathology 58: 100-110.

Motisi, N., Doré, T., Lucas, P., Montfort, F. 2010. Dealing with the variability in biofumigation efficacy through an epidemiological framework. Soil Biology and Biochemistry 42, 2044-57.

Murray, D. I. L. 1981. Rhizoctoina solani causing barley stunt disorder. Transactions of
the British Mycological Society. 76: 383–395.

Mutch, D. R., and Snapp, S. S. 2003. Cover crop choices for Michigan. Michigan State University Extension.

Nelson, E. B., and Hoitink, H. A. J. 1982. Factors affecting suppression of Rhizoctonia solani in container media. Phytopathology, 72(3), 275-279.

Ogoshi, A. 1987. Ecology and pathogenicity of anastomosis and intraspecific groups of
Rhizoctonia solani Kuhn. Ann. Rev. Phytopathol. 25: 125-143.

Pane, A., Martini, P., Chimento, A., Rapetti, S., Savona, S., Grasso, F. M., Cacciola, S. O., 2005. Phytophthora species on ornamental plants in Italy. Journal Of Plant Pathology 87: 301 (Abstract).

Peace, T. R. 1962. Pathology of trees and shrubs. Oxford University Press. 753 p.

Pierson, V. G. 1959. Methods of testing sugar beets to Rhizoctonia solani. Colorado State University – Master of Science Thesis.  pp. 75, 2959.

Richards, T. A., Dacks, J. B., Jenkinson, J. M., Thornton, C. R. and Talbot, N. J. 2006. Evolution of filamentous plant pathogens: gene exchange across eukaryotic kingdoms. Current Biology 16: 1857–1864.

Rosen, D. 1990. Biological control: selected case histories. Armored scale insects. Their biology, natural enemies and control. Vol. 4B. Elsevier Amsterdam, 497-505.

Sang, J. P., Minchinton, I. R., Johnstone, P. K., and Truscott, R. J. W. 1984. Glucosinolate profiles in the seed, root and leaf tissue of cabbage, mustard, rapeseed, radish and swede. Can. J. Plant Sci. 64, 77-93.

Sippell, D. W., Davidson, J. G. N., Sadasivaiah, R. S. 1985. Rhizoctonia root rot of rapeseed in the Peace River region of Alberta. Can. J. Plant Pathol. 7: 184–186.

Snapp, S., Date, K., Cichy, K., and O’Neil, K. 2006. Mustards–a Brassica cover crop for Michigan. Extension Bulletin E-2956, Michigan State University Extension, USA.

Spencer, D. and Fox, R. A. 1978. Assessment of pathogenicity of Rhizoctonia solani Kühn to some potato tissues and to cereals. Potato Research. 21: 81–88.

Stahl, B. 2004. Theophrastaceae. Pp. 472-478 in K. Kubitzki (ed.), The families and genera of vascular plants. Vol. VI. Flowering plants. Dicotyledons. Celastrales, Oxalidales, Rosales, Cornales, Ericales. Springer-Verlag, Berlin.

Stephens, C. T., Herr, L. F., Schmitthenner, A. F., Powell, C. C. 1982. Characterization of Rhizoctonia isolates associated with damping-off of bedding plants. Plant Dis. 66: 700–703.

Sundermeier, A. 2008. Oilseed radish cover crop. Ohio State Univ. Extension Fact Sheet SAG-5-08, Columbus.

Tahvonen, R., Hollo, J., Hannukkala, K. A. 1984. Rhizoctonia solani damping-off on spring turnip rape and spring rape (Brassica spp.) in Finland. J. Agric. Sci. Finland 56: 143–15.

Weller, D. M., Cook, R. J., MacNish, G. C., Basset, E. N., Powelson, R. L and Peterson, R. R. 1986. Rhizoctonia root rot of small grains favored by reduced tillage in the Pacific Northwest. Plant Disease. 70: 70–73.

Yang, J., Verma, P. R., Tewari, J. P. 1992. Histopathology of resistant mustard and susceptible canola hypocotyls infected by Rhizoctonia solani. Mycol. Res. 96 171–179. 10.1016/S0953-7562 (09)80962-3. 

TABLES AND FIGURES

Table 3. Effect of Rhizoctonia solani inoculation methods and inoculum levels on root rot disease severity percentages of Viburnum odoratissimum ‘sweet viburnum’ plants

Inoculation method

Inoculum level

Root rot disease severity (%)zyz

Plant fresh weight (g)xz

Agar slurry

3 petri dishes/L

69.3 a

7.0 c

Agar slurry

2 petri dishes/L

66.1 a

6.6 c

Agar slurry

1 petri dishes/L

63.0 a

8.4 c

Agar plugs

1 plug

31.8 b

8.8 c

Agar plugs

3 plugs

38.0 b

9.4 abc

Agar plugs

5 plugs

41.1 b

9.4 abc

Chopped potato soil medium

0.5 g/Kg-1 mix

31.8 b

9.6 abc

Chopped potato soil medium

1 g/Kg-1 mix

34.9 b

12.4 ab

Chopped potato soil medium

1.5 g/Kg-1 mix

38.0 b

12.9 a

Non-inoculated control

11.3 c

12.4 ab

P-value 

0.000

0.003

zRoot disease severity values of each represent the final visual ratings (0-5) converted into percentages where; 1(0%) = no symptom, heavily branched root system, and healthy looking; 2 (1-25%) = light brown necrosis in distinct spots, often necrosis in the root tip, less branched root system than healthy roots; 3 (26-50%) = few side roots, and dark brown necrosis in distinct spots; 4 (51-75%) = few and small side-roots, and dark brown necrosis of most of the root system, or all around the stem; 5 (76-100%) = plant dead. 

yMeans that do not share a letter are significantly different, according to the Fisher’s protected least significant difference test.

xPlant fresh weight were measured in grams at the end of the experiment.

z Experiment was repeated twice and values represent the mean values of the two experiments, with six replications per experiment.

Table 4. Root rot disease severity and disease incidence of tested Brassicaceae cover crops when inoculated with Rhizoctonia solani and Phytophthora nicotianae in greenhouse experiments

Cover crop

Root rot disease severity (%)zy

Disease incidence 

(%)x

R. solani

P. nicotianae

R. solani

P. nicotianae

Amara mustard green

7.7 e

23.4 d

1.9 d

36.9 c

Arugula

77.6 abc

44.3 c

66.7 ab

26.9 cd

Astro arugula

19.3de

19.3 def

1.2 d

13.5 cde

Bellezia arugula

85.9 a

77.9 a

66.7 ab

90.7 ab

Dragon’s tongue arugula

71.3 bc

54.7 bc

83.3 ab

100.0 a

Dwarf essex rape Brassica

15.1de

10.8 d-g

1.1 d

9.5 de

Mighty mustard® pacific gold

23.4 d

13.5 cde

0.8 d

11.7 cde

Mustard

21.3 d

9.8 efg

17.9 cd

18.5 cde

Oilseed radish

25.5 d

1.1 g

0.9 d

0.0 e

Olive-leaved sylvetta arugula

81.8 ab

65.1 ab

100 a

86.4 ab

Oriental mustard

15.1 de

21.2 de

0.7 d

34.9 cd

Purple top forage turnips

19.3 de

7.7f g

0.9 d

13.3 cde

Red giant mustard green

73.4 bc

60.9 b

16.8 cd

90.6 ab

Sylvetta green arugula

67.2 c

54.7 bc

50.0 bc

65.2 d

Wasabi arugula

77.6 abc

54.7 bc

50.0 bc

88.9 ab

P– value

0.000

0.000

0.000

0.000

zRoot rot disease severity values of each represent the final visual ratings (0-5) converted into percentages where; 1(0%) = no symptom, heavily branched root system, and healthy looking; 2 (1-25%) = light brown necrosis in distinct spots, often necrosis in the root tip, less branched root system than healthy roots; 3 (26-50%) = few side roots, and dark brown necrosis in distinct spots; 4 (51-75%) = few and small side-roots, and dark brown necrosis of most of the root system, or all around the stem; 5 (76-100%) = plant dead. 

yValues represent the means of two experiments, with six replications per experiment. Disease symptoms were not observed on or nor was recovered from the control plants. Means that do not share a letter are significantly different, according to the Fisher’s protected least significant difference test.

xDisease incidence values represent the percentage of plants which shown disease symptoms throughout the experiment, compared to the germination percentages of the control plants. Symptoms recorded include pre- and post-damping off, chlorosis, wilting and plant death.

Table 5. Pearson correlation between plant growth data and root rot disease severity and disease incidence for all cultivars caused by Rhizoctonia solani and Phytophthora nicotianae in greenhouse trials 

Growth data

Root rot disease severity (%)z

Disease incidence (%)z

R. solani

P. nicotianae

R. solani

P. nicotianae

Plant height

R2= -0.782

P= 0.00

R2= -0.811

P= 0.00

R2= -0.712

P= 0.00

R2= -0.798

P= 0.00

Plant width

R2= -0.785

P= 0.00

R2= -0.823

P= 0.00

R2= -0.665

P= 0.00

R2= -0.807

P= 0.00

Plant fresh weight

R2= -0.751

P= 0.00

R2= -0.777

P= 0.00

R2= -0.661

P= 0.00

R2= -0.731

P= 0.00

zNegative values represent a negative correlation and values closer +1 or -1 indicates a strong relationship between growth data and root rot disease severity and disease incidence. Values represent the means of two experiments, with six replications per experiment and root rot disease severity and incidence for all cover crops. 

Table 6. Effect of Rhizoctonia solani and Phytophthora nicotianae on plant height and fresh weight reduction of selected Brassicaceae cover crops in greenhouse pathogenicity experiment

Cover crop

Reduction in plant height (%)zyw

Reduction in plant fresh weight (%)zxw

R. solani

P. nicotianae

R. solani

P. nicotianae

Amara mustard green

3.5 f

17.9 a

13.5 bcd

16.3 a

Arugula

6.2 ef

12.8 a

6.6 cd

3.6 a

Astro arugula

12.6 c-f

9.0 a

9.4 bcd

4.2 a

Bellezia arugula

15.1 b-f

21.8 a

44.5 a

28.6 a

Dragon’s tongue arugula

26.2 ab

27.3 a

29.0 abc

32.0 a

Dwarf essex rape Brassica

22.0 abc

2.7 a

32.5ab

8.4 a

Mighty mustard® pacific gold

8.4 def

4.7 a

17.8 bcd

13.3 a

Mustard

19.6 a-d

11.6 a

4.1 d

10.5 a

Oilseed radish

8.8 def

3.3 a

13.5 bcd

3.0 a

Olive-leaved sylvetta arugula

10.5 c-f

20.9 a

29.9 abc

23.5 a

Oriental mustard

9.3 c-f

13.5 a

24.9 a-d

18.3 a

Purple top forage turnips

16.7 a-d

4.9 a

2.4 d

7.6 a

Red giant mustard green

26.3 ab

9.6 a

30.7 abc

23.6 a

Sylvetta green arugula

30.0 a

27.3a

32.3 ab

28.9a

Wasabi arugula

11.4 a-d

13.9 a

21.5 a-d

22.6 a

P– value

0.003

0.248 

0.037

0.059 

zValues represent the mean values of the two experiments, with six replications per experiment. Means that do not share a letter are significantly different at P0.059, according to the Fisher’s protected least significant difference test.

yPlant height was measured in centimeters at the end of the experiment.

xPlant fresh weight were measured in grams at the end of the experiment.

wReduction in plant height and fresh weight (%) for each pathogen compared with control plants.

Figure 1. Effect of Rhizoctonia solani and Phytophthora nicotianae on fresh weight of four Brassicaceae crops grown commercially for biofumigation. Each column represents means of two experiments, with six replicate plants per pathogen per experiment. Columns with a letter in common are significantly different within each of the cover crop based on Fisher’s protected least significant difference test. 

Figure 2. Effect of Rhizoctonia solani and Phytophthora nicotianae on root rot disease severity of selected Brassicaceae crops. The plants showed significantly higher root rot disease percentages when inoculated with R. solani compered to P. nicotianae (P<0.05). Each column represents means of two experiments, with six replicate plants per pathogen per experiment. Columns with a letter in common are significantly different within each of crop based on Fisher’s protected least significant difference test.

Objective 2. Determining the effect of biofumigation in suppressing Rhizoctonia solani and Phytophthora nicotianae in woody ornamental nurseries

RESULTS

Identifying volatile compounds in soils biofumigated with cover crops. Different types of ITC compounds and derivatives of ITC (nitriles, thiocyanate, thiol, amide) were identified from the GC-MS results (Table 2). Allyl isothiocyanate was observed from mighty mustard® pacific gold, amara mustard green and oriental mustard at RT = 5.2-5.3 min retention time when sampled 24- hrs after the incorporation; same compound was detected from amara mustard also 7 days after soil incorporation. Benzyl isothiocyanate was recorded from mighty mustard® pacific gold, dwarf essex rape and mustard (S. alba) at RT = 14.40 min.  3-Butenyl isothiocyanate and 4-methylthio-3 butenyl isothiocyanate were detected 24-hrs after the incorporation of purple top forage turnips and radish, respectively. Benzyl isothiocyanate was only detected 24-hrs after the amendment of mustard (S. alba).

Effect of biofumigation in suppressing Rhizoctonia solani. All the eight Brassicaceae cover crops significantly reduced disease severity caused by R. solani on viburnum roots when compared to the inoculated, non-biofumigated control (Table 3). Biofumigation time did not affect disease severity, as similar disease severity values were recorded from both 2 and 4 weeks Brassica cover cropped experiments. In both experiments the inoculated, non-biofumigated control plants had significantly greater root rot disease severity and the non-inoculated, non-biofumigated control plants displayed the lowest disease severity levels. Whether biofumigation was performed over a 2 weeks or 4 weeks interval, viburnum plant roots from amara mustard green cover crop-incorporated pots had the lowest Rhizoctonia root rot disease severity, followed by the mighty mustard® pacific gold cover crop.

The highest Rhizoctonia pathogen recoveries from roots were recorded from the inoculated, non-biofumigated controls (100%) and radish amended soil (100%) (Table 3). Less recovery was recorded from mustard (75%)-incorporated pots at 4 weeks biofumigation but values were not statistically different than other cover crops except for radish. Amara mustard green and astro arugula had numerically lower pathogen recovery percentages (60%, respectively) but were not statistically different from other Brassica cover crops except for radish- and dwarf essex rape-incorporated pots in the 2 weeks biofumigation experiment (Table 3). Among the tested cover crops; amara mustard green, astro arugula, purple top forage turnips, mighty mustard® pacific gold and mustard had lowest Rhizoctonia root rot disease percentages and pathogen recovery from the roots in both experiments.

In the cover crop treated pots of both experiments, the highest number of Rhizoctonia colony forming units (CFU) log values in R. solani selective medium were recorded just prior to incorporation of cover crops to the pots (Table 4). Rhizoctonia colony quantities began decreasing after 24-hrs of biofumigation, with continuous reduction over the biofumigation time interval in both experiments, except for radish at 4 weeks after biofumigation that showed increased CFU values. Prior to the cover crop incorporation, initial fungal populations in the R. solani selective medium of all the treatments were similar to inoculated, non-biofumigated control. Twenty-four hours after cover crop incorporation, the highest fungal populations were recorded in inoculated, non-biofumigated controls (log values 3.89, 3.94) at a statistically significant rate compared to all cover crops of both 2 and 4 weeks biofumigation experiments, respectively. From the 24-hrs sample onward, all Brassica cover crops-incorporated soils in both experiments showed significantly lower fungal populations compared to the inoculated, non-biofumigated controls, but significantly higher colony counts than non-inoculated, non-biofumigated controls (Table 4). Randomly selected colonies were re-cultured on R. solani selective medium and identified as R. solani using DNA sequencing.

Initial heights of the viburnum cuttings ranged from 19.00 to 31.00 cm for the 4 weeks experiment, and 12.00 to 18.00 cm for the 2 weeks experiment (data not shown). Due to this high variation in initial cutting height, final heights were also measured and the change in height calculated at the end of the experiments. In the 4 weeks biofumigation experiment, the greatest increase in height was observed in plants grown in amara mustard green (14.67 cm)-incorporated pots, followed by the astro arugula (14.33 cm), then purple top forage turnips (8.67 cm), but these differences were not statistically significant (Table 5). Viburnum plants grown in these three cover-crop-incorporated pots had significantly higher increases than the inoculated, non-biofumigated controls or other cover crops, but they did not show a statistically significant height increase when compared with the non-inoculated, non-biofumigated control plants. Astro arugula (8.00 cm), amara mustard green (7.00 cm), dwarf essex rape (7.33 cm) and mustard  (6.33 cm) showed the greatest height increases when compared to the inoculated, non-biofumigated control plants and the other cover crops in the 2 weeks experiment (Table 5).

Change in plant width (P = 0.009) was also recorded at the end of the each experiment. Viburnum plants grown in astro arugula-incorporated pots had the greatest width increase (21.67 cm), followed by mighty mustard® pacific gold (20.33 cm), purple top forage turnips (15.67 cm) and amara mustard green (15.33 cm), but the width increases were not significantly different from the non-inoculated, non-biofumigated control pots (15.17 cm). In the 2 weeks biofumigation experiment, width increase ranged from 1.33 to 7.16 cm, which was less than the 4 weeks biofumigation experiment (6.67 cm- 21.67 cm). The greatest width increments were recorded by plants grown on amara mustard green (7.17 cm), purple top forage turnips (6.33 cm) and astro arugula (5.83 cm)-incorporated pots (Table 5). Plants were sustained for nearly 4 months after biofumigation in the 4 weeks experiment and for nearly 2-months after biofumigation in the 2 weeks experiment before termination. Extended time period can be the possible reason for this drastic difference in the width increment between the two experiments. 

A significant negative correlation was observed between root rot percentage and plant fresh weight in both the 4 weeks biofumigation experiment (R2 = -0.612, P = 0.000) and 2 weeks biofumigation experiment (R2 = -0.581, P = 0.001) (data not shown). In both experiments, the highest plant fresh weight was recorded by plants grown in amara mustard green- or astro arugula-incorporated soil. Plant fresh weights were numerically different compared to non-inoculated, non-biofumigated pots, but statistically different from inoculated, non-biofumigated pots (Table 6). Viburnum plants grown in oriental mustard- or radish-incorporated soil demonstrated the second-lowest fresh weight, higher only than the inoculated, non-biofumigated controls in both of the experiments. The amara mustard green- and mighty mustard® pacific gold-incorporated soils significantly increased the root weight of viburnum plants compared to other cover crops and the non-biofumigated controls in the 4 weeks biofumigation experiment. Astro arugula, mustard and dwarf essex rape-incorporated soils significantly increased root weight of viburnum plants compared to other cover crops and the non-biofumigated controls in 2 weeks experiment.

Effect of biofumigation in suppressing Phytophthora nicotianae. All the tested Brassica cover crops except radish were effective in reducing disease severity caused by P. nicotianae on hydrangea roots in the 4 weeks biofumigation experiment when compared to the inoculated, non-biofumigated control (Table 7). Phytophthora root rot disease severity was between 8.67 – 69.25% in the 4 weeks biofumigation experiment and 2.25 – 65.08% in the 2 weeks experiment. In both biofumigation experiments, the highest disease severity was recorded by the inoculated, non-biofumigated controls, while least disease severity occurred in non-inoculated, non-biofumigated control plants. In the 4 weeks biofumigation experiment, astro arugula and dwarf essex rape were the most effective cover crops at reducing Phytophthora root rot severity. In the 2 weeks biofumigation experiment, amara mustard green, astro arugula and dwarf essex rape were the most effective cover crops at reducing Phytophthora root rot severity. In both experiments, hydrangea plants grown in astro arugula-, dwarf essex rape- and amara mustard green-incorporated soil had low pathogen recovery from cultured roots (Table 7). In both experiments, significantly fewer amounts of Phytophthora colony forming units on PARPH-V8 medium were recorded in the non-inoculated, non-biofumigated controls before incorporation of the cover crops (Table 8). Twenty-four hours after the incorporation of the cover crops, all the treatments except radish had significantly lower CFU log values compared to the inoculated, non-biofumigated control. In both experiments, CFU log values of radish- and turnip-incorporated soil were not significantly different from the inoculated, non-biofumigated control, while all other cover crop amended treatments had significantly lower CFU values. Astro arugula-, amara mustard green-, mustard (S. alba)- and dwarf essex rape-incorporated soil samples had significantly lower CFU values four weeks after biofumigation in the 4 weeks experiment, and there were no significant differences in the CFU values between these cover crops incorporated soils and the non-inoculated, non-biofumigated control. At the end of the 4 weeks biofumigation experiment, the CFU values for all cover crop incorporated soils increased to values significantly higher than the non-inoculated, non-biofumigated control, but statistically lower than the inoculated, non-biofumigated control soil. In the 2 weeks biofumigation experiment, soil samples taken from astro arugula- and mustard (S. alba)-incorporated pots had the lowest log value of Phytophthora compared to the other cover crops, and there were no significant differences in the CFU values between these two cover cropped soils and the non-inoculated, non-biofumigated control soil. Ten weeks after biofumigation, amara mustard green- and dwarf essex rape-incorporated soil samples had the lowest log value of Phytophthora compared to the other cover crops. 

Hydrangea rooted cuttings used in both experiments had similar heights at the beginning of the experiments (P = 0.356 and P = 0.162) (data not shown). Similar observations were made regarding plant width data in both experiments. Plant width observations taken at the beginning and end of the each experiment and overall width increase was not significantly different from the widths recorded in the inoculated control plants (data not shown). Greatest width increase was recorded by hydrangea rooted cuttings grown on amara mustard green-incorporated pots (Table 9). 

Hydrangea plants grown in astro arugula-, dwarf essex rape- and amara mustard green-incorporated soils had significantly higher plant fresh weights and root weights when compared to the inoculated, non-biofumigated control plants and the other cover crops in the 4 weeks experiment. There were no significant differences among these cover crops and non-inoculated, non-biofumigated control in either whole plant weight or root weight (Table 10). In the 2 weeks experiment, astro arugula-incorporated soil was most effective in increasing hydrangea plant weight.  Astro arugula-, amara mustard green-, dwarf essex rape- and purple top forage turnip-incorporated soils were equally effective in increasing hydrangea root weight. Phytophthora root rot disease severity had a significantly negative effect on plant fresh weight in both experiments (4 weeks experiment R2 =-0.562, P = 0.001; 2 weeks experiment R2 =- 0.576, P = 0.001) (data not shown). 

Effect of biofumigation on soil chemistry. All soil samples taken from the tested pots (2 weeks and 4 weeks experiments) had soil pH mean values in between 5.43 – 6.08. Soil analysis showed purple top forage turnip- (6.08), oriental mustard- (6.01) and dwarf essex rape- (6.00) incorporated soil had the highest soil pH values, while radish- (5.43), mighty mustard® pacific gold- (5.78) incorporated and the inoculated, non-biofumigated control (5.60) soils had the lowest (data not shown). Soil phosphorus (P) values ranged from 9 lbs/A (mustard (S. alba), oriental mustard and non-inoculated, non-biofumigated control) to 14 lbs/A (radish). Radish also had the highest amount of soil potassium (K) at a rate of 123 lbs/A, while the non-inoculated, non-biofumigated controls (47 lb/A) had the lowest. Soil calcium (Ca) levels were highest in purple top forage turnip-incorporated (1,620 lb/A), the inoculated, non-biofumigated controls (1,572 lb/A), and astro arugula-incorporated (1,544 lb/A) soil, and lowest in radish-incorporated soil (1,119 lb/A) and the non-inoculated, non-biofumigated controls (1,205 lb/A). Purple top forage turnip-incorporated soil had the highest magnesium, iron and manganese, while samples taken from the non-inoculated, non-biofumigated control pots had the lowest. Boron levels were constant at 0.3 lbs/A in all treatments apart from 0.4 lbs/A in purple top forage turnip-incorporated soil and the non-inoculated, non-biofumigated control soils. Soil nitrate content was highest in astro arugula- (35.14 ppm) and radish- (15.74 ppm) incorporated soil samples and lowest in mighty mustard® pacific gold- (1.36 ppm) and oriental mustard- (1.52 ppm) incorporated soil. Soil organic matter percentage ranged from 1.9% (astro arugula) to 1.36% (amara mustard green and non-inoculated, non-biofumigated soil).

DISCUSSION   

Soil fumigants have been widely used in agriculture to control plant diseases and achieve higher crop yields. Methyl bromide and chloropicrin were historically the most commonly used fumigants to control fungal pathogens (Lazzeri et al. 2003) but are detrimental to the environment. Metam sodium is another fumigant used to control soilborne pathogens (Larkin and Griffin 2006), but it decreases microbial biomass and activity, negatively affects microbial community structure (Omirou et al. 2011). It is therefore important to find alternative, environmentally friendly approaches to control soilborne diseases. 

R. solani and P. nicotianae disease suppression from biofumigation may not be primarily caused by the presence of cyanate compounds but could be due to other GSL hydrolysis products or biotic factors. To collect AITC, volatiles need to be collected within 24 hrs of biofumigation, while other GSL hydrolysis products could be detected in the soil up to 14-days post-biofumigation. The process of collecting volatiles directly from the treatment pots was challenging due to the high moisture content of the media during the fumigation process. In future studies, we recommend that separate soil samples be used for volatile studies so the media can be manipulated to maximize volatile collection efficacy. We also suggest analyzing freeze-dried and freshly macerated plant material in order to distinguish volatiles released directly from the plants from those subsequently modified within the soil. LC-MS analysis of glucosinolate precursors would also provide additional information regarding potential volatile metabolites. 

Soil type and soil environment affects ITC release and can be a possible reason for the variations observed among related similar studies. Higher clay or peat content reduces the efficacy of ITC (Brown and Morra 1996; Matthiessen and Shackleton 2005). Bending and Lincoln (2000) found gas phase AITC concentrations were lower in clay loam soils than in sandy loam soil due to the adsorption of AITC by organic carbon. Adsorption of AITC into soil has been shown to increase with an increase in organic matter content (Price 2005). In our study, the topsoil had high clay content, which could have contributed to low detection of ITC compounds.

Phytotoxicity is one of the important factors when selecting biofumigant cover crops. While biofumigation can suppress pathogen, if the incorporated plant material negatively affects growth of the subsequent crop as well it will not be economically feasible. Lazzeri et al. (2009) did not observe phytotoxicity symptoms in transplanted zucchini plants after mustard meal incorporation. In our study, phytotoxicity was not observed on viburnum and hydrangea woody ornamental plants after either the 14-day or 1-month biofumigation period with any of the tested cover crop. 

In this research, mustard (S. alba), purple top forage turnips (B. rapa), astro arugula (E. vesicaria spp. sativa), mighty mustard® pacific gold (B. juncea), dwarf essex rape (B. napus), amara mustard green (B. carinata) and oriental mustard (B. juncea) cover crops were effective in controlling R. solani and P. nicotianae pathogens in nursery soils. Similar disease suppression was observed whether biofumigation was performed for 1-month or 14-days, suggesting that the increased biofumigation intervals may not increase the biofumigation effectiveness.

Viburnum odoratissimum plants were highly susceptible to R. solani and Hydrangea paniculata plants also showed root rot symptoms due to P. nicotianae infection. Both R. solani and P. nicotianae pathogens negatively affected the plant fresh weights of viburnum and hydrangea plants, however, viburnum and hydrangea grown in amara mustard green-, astro arugula- and purple top forage turnip-incorporated soil had significantly higher plant and root fresh weight compared to the inoculated, non-biofumigated control plants.  

 To date, more than 20 aromatic and aliphatic isothiosyanates and other potential allelochemicals have been identified from Brassica juncea, B. napus, B. campestris and B. nigra (Brown and Morra 1995 and 1996). Depending on the GSL profile, each Brassica species releases different cyanates. Allyl isothiocyanate (AITC) has a suppressive effect on R. solani and P. ultimum. In a controlled laboratory study, it had a fungicidal effect on R. solani at a rate of 2.2 µmol L-1 and 3.3 µmol L-1, and reduced growth of the pathogen by 27% and 55%, respectively (Charron et al. 1999). Although methyl isothiocyanate (MITC) is the simplest structure of the many ITCs that may be produced through Brassica tissue incorporation, it is not a commonly found ITC in Brassicaceae crops (Matthiessen and Shackleton 2005). The most available and studied ITC is AITC, which is produced from allyl GSL. Allyl GSL is commonly found in B. juncea, B. napus, and B. carinata, but many Brassica cover crops can produce AITC through hydrolysis of GSL at a pH of 4.0 or greater (Mayton et al. 1996). In this study, we were able to identify AITC in amara mustard green (B. carinata), mighty mustard® pacific gold (B. juncea), and oriental mustard (B. juncea) while soil pH was 5.72, 5.78 and 6.01, respectively. While amara mustard green and mighty mustard® pacific gold were effective in controlling R. solani, oriental mustard was less effective.

Although amara mustard green and astro arugula are not currently used as commercial biofumigation cover crops, according to our study they show promising results in controlling soilborne pathogens of woody ornamental plants in greenhouse conditions. Further experiments should be conducted to test the ability of these Brassica cover crops to control other major pathogens in woody ornamental nursery production and in other agricultural production systems.  

LITERATURE CITED

Ali, J. G., Alborn, H. T., Campos-Herrera, R., Kaplan, F., Duncan, L. W., Rodriguez-Saona, C., and Stelinski, L. L. 2012. Subterranean, herbivore-induced plant volatile increases biological control activity of multiple beneficial nematode species in distinct habitats. PLoS One, 7(6), e38146.

Banks, J. G., Board, R. G., Sparks, N. H. C. 1986. Natural antimicrobial systems and their potential in food preservation of the future. Biotechnology Applied Biochemistry 8: 103-107.

Bellostas, N., Casanova, E., Garcia-Mina, J. M., Hansen, L. M., Jørgensen, L. N., Kudsk, P., Madsen, P. H., Sørensen, J. C. and Sørensen, H. 2007. Biological activity of glucosinolate derived compounds isolated from seed meal of Brassica crops and evaluated as plant and food protection agents. Organic & Prints.

Benderoth, M., Textor, S., Windsor, A. J., Mitchell-Olds, T., Gershenzon, J., and Kroymann, J. 2006. Positive selection driving diversification in plant secondary metabolism. Proceedings of the National Academy of Sciences, 103(24), 9118-9123.

Bending, G. D., Lincoln, S. D. 2000. Inhibition of soil nitrifying bacteria communities and their activities by glucosinolate hydrolysis productsSoil Biol. Biochem. 32, 1261–1269 10.1016/S0038-0717(00)00043-2.

Björkman, R. 1973. Interaction between proteins and glucosinolate isothiocyanates and oxazolidinethiones from Brassica napus seed. Phytochemistry, 12, 1585-1590.

Blok, W. J., Lamers, J. G., Termorshuizen, A. J., Bollen, G. J. 2001. Control of soilborne plant pathogens by incorporating fresh organic amendments followed by tarping. Phytopathology 90: 253-259.

Borek, V., Morra, M. J., Brown, P. J., McCaffery, J. P. 1995. Transformation of the glucosinolate-derived allelochemicals allyl isothiocyanate and allylnitrile in soil. J Agric Food Chem 43:1935–1940.

Brown, P. D., and Morra, M. J. 1995. Glucosinolate-containing plant tissues as bioherbicides. J. Agric. Food Chem. 43:3070-3074.  

Brown, P. D., and Morra, M. J. 1996. Hydrolysis products of glucosinolates in Brassica napus tissues as inhibitors of seed germination. Plant Soil 181:307-316.

Ceustermans, A., Van Wambeke, E., Coosemans, J. 2010. Efficacy of chemical alternatives for methyl bromidein lettuce e production: field experiment. Acta Hort. (ISHS) 883:135–143.

Charron, C. S., and Sams, C. E. 1999. Inhibition of Pythium ultimum and Rhizoctonia solani by shredded leaves of Brassica species. Journal of the American Society of Horticultural Science 124(5): 462–467.

Dungan, R. S., Gan, J., Yates, S. R., 2003. Accelerated degradation of methyl isothiocyanate in soil. Water, Air, and Soil Pollution 142, 299–310.

Dunne, C. 2002. Comparison of integrated management practices for the control of Phytophthora cinnamomi in protea cultivation in WA, Floriculture News, no. 56, March.

Fahey, J. W., Zalcmann, A. T., and Talalay, P. 2001. The chemical diversity and distribution of Glucosinolates and Isothiocyanates among plants. Phytochemistry, 56, 5-51.

Falk, A., Xue, J., Lenman, M., and Rask, L. 1992. Sequence of a cDNA clone encoding the enzyme myrosinase and expression of myrosinase in different tissues of Brassica napus. Plant Sci. 83, 181-186.

Ferguson, A. J., and Jeffers, S. N. 1999. Detecting multiple species of Phytophthora in container mixes from ornamental crop nurseries. Plant Dis. 83(12):1129-1136.

Gardiner, J. B., Morra, M. J., Eberlein, C. V., Brown, P. D., Borek, V., 1999. Allelochemicals released in soil following incorporation of rapeseed (Brassica napus) green manures. Journal of Agriculture and Food Chemistry.

Gilardi, G. Gullino, M. L.Garibaldi, A. 2013. Critical aspects of grafting as a possible strategy to manage soil-borne pathogens. Scientia Horticulturae; 149:19-21.

Gimsing, A., Kirkegaard, J. 2009. Glucosinolates and biofumigation: fate of glucosinolates and their hydrolysis products in soil. Phytochemistry Reviews 8, 299-310.

Gimsing, A.L., Kirkegaard, J.A. 2006. Glucosinolate and isothiocyanate concentration in soil following incorporation of Brassica biofumigants. Soil Biol Biochem 38:2255–2264.

Goud, T. Y., Devi, G. U., Reddy, P. N., and Sankar, A. S. 2013. Activity of volatile toxins of brassica residues on stem and pod rot disease of groundnut caused by S. rolfsii under greenhouse conditions. Annals of Biological Research, 4(9), 63-66.

Gould, W. D., Hagedorn, C., Bardinelli, T. R., and Zablotowicz, R. M. 1985. New selective media for enumeration and recovery of fluorescent Pseudomonads from various habitats. Applied and Environmental Microbiology, 49(1), 28-32.

Gutierrez, W. A., Shew, H. D., Melton, T. A. 2001. A semi-selective medium to isolate Rhizoctonia solani from soil and tissue. Plant Pathological Extension, 1-2.

Henry, M., Béguin, M., Requier, F., Rollin, O., Odoux, J. F., Aupinel, P. 2012. Response to comment on “a common pesticide decreases foraging success and survival in honey bees.” Science 337, 1453. doi: 10.1126/science. 1224930.

Hollister, E. B., Hu, P., Wang, A. S., Hons, F. M., and Gentry, T. J. 2013. Differential impacts of brassicaceous and nonbrassicaceous oilseed meals on soil bacterial and fungal communities. FEMS Microbiol. Ecol. 83, 632–641. doi: 10.1111/1574-6941.12020.

Holmes, K. A., and Benson, D. M. 1994. Evaluation of Phytophthora parasitica var. nicotianae for biocontrol of Phytophthora parasitica on Catharanthus roseus. Plant disease, 78(2), 193-199.

Howell, C. R., Stipanovic, R. D. 1979. Control of Rhizoctonia solani on cotton seedlings with Pseudomonas fluorescens and with an antibiotic produced by the bacterium. Phytopathology, 69, 480-482. 

Kawakishi, S., Kaneko, T. 1985. Interaction of oxidized glutathione with allyl isothiocyanatePhytochemistry 24, 715–718 10.1016/S0031-9422(00)84882-7.

Kelly, P. J., Bones, A. and Rossiter, J. T. 1998. Sub-cellular immunolocalization of the glucosinolate sinigrin in seedlings of Brassica juncea.  Planta 206: 370–377.

Kirkegaard, J. A., and Matthiessen, J. N. 2005. Developing and refining the biofumigation concept. AgroIndustria, 3, 5-11.

Kojima, M., Oawa, K. 1971. Studies on the effect of isothiocyanates and their analogues on microorganisms. (I) Effects of isothiocyanates on the oxygen uptake of yeasts. Journal of Fermentation Technology, 49, 740–746.

Koron, D., Sonjak, S., Marjana Regvar, M. 2014. Effects of non-chemical soil fumigant treatments on root colonization with arbuscular mycorrhizal fungi and strawberry fruit production. Crop Prot., 55, 35–41.

Larkin, R. P., and Griffin, T. S. 2006. Control of soilborne potato diseases using brassica green manures. Crop Protection 26: 1067-107.

Lazzeri, L., Baruzzi, G., Malaguti, L., and Antoniacci, L. 2003. Replacing methyl bromide in annual strawberry production with glucosinolate-containing green manure crops. Pest Manage. Sci. 59:983-990.

Lazzeri, L., D’Avino, L., Mazzoncini, M., Antichi, D., Mosca, G., Zanetti, F.,  and Cosentino, S. 2009. On-farm agronomic and first environmental evaluation of oil crops for sustainable bioenergy chains. Italian Journal of Agronomy, 4(4), 171-180.

Lazzeri, L., Malaguti, L., Cinti, S. 2013. The Brassicaceae biofumigation system for plant cultivation and defence. An Italian twenty-year experience of study and application. Acta Hort. (ISHS) 1005, 375-82.

Liyanapathiranage, P. 2017. Sustainable management of soilborne diseases in nursery production. MS Thesis. Tennesse State University? Nashville, TN.

Matthiessen, J. N., Kirkegaard, J. A. 2006. Biofumigation and enhanced biodegradation: opportunity and challenge in soilborne pest and disease management. Critical Reviews in Plant Sciences 25, 235-65.

Matthiessen, J. N., Shackleton, M. A. 2005. Biofumigation: environmental impacts on the biological activity of diverse pure and plant-derived isothiocyanates. Pest Manag. Sci. 61, 1043–1051. 10.1002/ps. 1086.

Mayton, H. S., Olivier, C., Vaughn, S. F., and Loria, R., 1996. Correlation of fungicidal activity of Brassica species with allyl isothiocyanate production in macerated leaf tissue. Phytopathology 86: 267-271.

Mazzola, M. 1998. Elucidation of the microbial complex having a causal role in the development of apple replant disease in Washington. Phytopathology 88:930–938.

Mazzola, M., Gu, Y.H. 2000. Impact of wheat cultivation on microbial communities from replant soils and apple growth in greenhouse trials. Phytopathology 90:114–119.

Mazzola, M., Brown, J., Izzo, A. D., and Cohen, M. F. 2007. Mechanism of action and efficacy of seed meal-induced pathogen suppression differ in a Brassicaceae species and time dependent manner. Phytopathology 97:454-460.

Morales-Rodríguez, C., Palo, C., Palo, E., Rodríguez-Molina, M.C. 2013. Control of Phytophthora nicotianae with mefenoxam, fresh Brassica tissues, and Brassica pellets. Plant Disease 98, 77-83.

Morra, M. J., and Kirkegaard, J. A. 2002. Isothiocyanate release from soil incorporated Brassica tissues. Soil Biol. Biochem. 34:1683-1690.

O’Neill, T. M., Rickwood, A., 2002. Project BOF45 – Final Report – Bulbs and cut flowers: development of a combined dazomet and metam-sodium treatment as an alternative to methyl bromide for soil sterilization. In. HDC.

Omirou, M., Rousidou, C., Bekris, F., Papadopoulou, K. K., Menkissoglou-Spiroudi, U., Ehaliotis, C. 2011. The impact of biofumigation and chemical fumigation methods on the structure and function of the soil microbial community. Microb. Ecol. 61, 201–213. 10.1007/s00248-010-9740-4. 

Poulsen, J. L., Gimsing, A. L., Halkier, B. A., Bjarnholt, N. and Hansen, H. C. B. 2008. Mineralization of benzyl glucosinolate and its hydrolysis product the biofumigant benzyl isothiocyanate in soil. Soil Biology and Biochemistry 40(1), 135–141.

Price, A. J. 2005. Allyl isothiocyanate and carbon dioxide produced during degradation of Brassica juncea tissue in different soil conditions, HortScience 40(6): 1734-1739.

Rosskopf, E. N., Chellemi, D. O., Kokalis-Burelle, N., and Church, G. T. 2005. Alternatives to methyl bromide: A Florida perspective. APSnet Feature, June.

Saharan, G. S., and Mehta, N. 2008. Economic importance. Sclerotinia diseases of crop plants: biology, ecology and disease management, 41-45.

Sang, J. P., Minchinton, I. R., Johnstone, P. K., and Truscott, R. J. W. 1984. Glucosinolate profiles in the seed, root and leaf tissue of cabbage, mustard, rapeseed, radish and swede. Can. J. Plant Sci. 64, 77-93.

Smolinska, U., Morra, M.J., Knudsen, G.R., James, R.L. 2003. Isothiocyanates produced by Brassicaceae species as inhibitors of Fusarium oxysporum. Plant Disease 87:407- 412.

Thangstad, O. P., Gilde, B., Chadchawan, S., Seem, M., Husebye, H., Bradley, D., and Bones, A. M. 2004. Cell specific, cross-species expression of myrosinases in Brassica napus, Arabidopsis thaliana and Nicotiana tabacum. Plant molecular biology, 54(4), 597-611.

Van Den Berg, F., Leistra, M., Roos, A. H., and Tuinstra, L. T. 1992. Sampling and analysis of the soil fumigants 1, 3-dichloropropene and methyl isothiocyanate in the air. Water, Air, and Soil Pollution, 61(3-4), 385-396.

 Van der Werf, H. M. 1996. Assessing the impact of pesticides on the environment. Agriculture, Ecosystems & Environment, 60(2-3), 81-96.

Xue, J., Pihlgren, U., and Rask, L. 1993. Temporal, cell-specific, and tissue-preferential expression of myrosinase genes during embryo and seedling development in Sinapis alba. Planta, 191(1), 95-101.

Yates, S. R., Ernst, F. F., Gan, J., Gao, F., & Yates, M. V. 1996. Methyl bromide emissions from a covered field: II. Volatilization. Journal of environmental quality, 25(1), 192-202.

Zsolnai, T. 1966. Antimicrobial effect of thiocyanates and isothiocyanates. Arztliche Forschung, 16, 870–876.

TABLES 

Table 1. Seeding rate of brassicaceae crops evaluated in this study

Brassicaceae crops

Scientific name

Number of seeds/A

Amara mustard green

Brassica carinata

2.3 x 106

Astro arugula

Eruca sativa

8.3 x 105

Dwarf essex rape 

B. napus

1.5 x 106

Mighty mustard® pacific gold

B. juncea

5.6 x 105

Mustard

Sinapis alba

2 x 106

Oriental mustard

B. juncea

5.6 x 105 

Purple top forage turnips

B. rapa

1.8 x 106 

Radish

Raphanus sativus

9.8 x 105

Table 2. Results from GC-MS analysis showing retention time, percent match and detection times of volatile compounds extracted from soils where different brassicaceae crops were incorporated

Brassicaceae crops

Compound

Retention time on DB 5 column 

(min)

Percent match to NIST08 database

Sampling time 

Amara mustard green

Allyl isothiocyanate 

5.30

84

24-hrs

5.31

88

7-days

Dwarf essex rape

Phenylethyl isothiocyanate 

14.41

93

24-hrs

Mighty mustard® pacific gold

Allyl isothiocyanate

5.29

86

24-hrs

Phenylethyl isothiocyanate 

14.40

94

7-days

Mustard

(S. alba)

Benzyl isothiocyanate 

13.06

94

24-hrs

Phenethyl isothiocyanate 

14.41

88

24-hrs

Oriental mustard

Allyl isothiocyanate 

5.29

70

24-hrs

Purple top forage turnips

3-Butenyl isothiocyanate 

7.05

86

24-hrs

Radish

4-methylthio-3butenyl isothiocyanate

14.00

84

24-hrs

Table 3. Rhizoctonia root rot disease severity on viburnum (Viburnum odoratissimum) and pathogen recovery from the roots grown in a soil biofumigated for 4 and 2 weeks

Treatment

Biofumigation 

(4 weeks)

Biofumigation 

(2 weeks)

Disease severity (%)z

Pathogen recovery from roots (%)xy

Disease severity (%)zx

Pathogen recovery from roots (%)xy

Amara mustard green        

15.08 efx

76.67 b

19.25 de

60.00 d

Astro arugula

27.58 cde

81.67 b

23.42 cd

60.00 d

Mighty mustard® pacific gold

19.25 e

82.50 b

23.42 cd

61.67 d

Mustard

35.92 cd

75.00 b

35.92 b

65.83 cd

Oriental mustard

40.08 bc

87.50 ab

40.08 b

67.50 cd

Radish

52.58 b

100.00 a

44.25 b

77.50 b

Dwarf essex rape

35.92 cd

86.67 b

40.08 b

89.17 ab

Purple top forage turnips

23.42 de

78.33 b

31.75 bcd

71.67 cd

Non-inoculated control  

4.50 f

21.67 c

6.75 e

15.00 e

Inoculated control

73.42 a

100.00 a

77.58 a

100.00 a

P-value 

<0.0001

<0.0001

<0.0001

<0.0001

 

zRoot rot disease severity was recorded at the end of the trial for each plant using a scale of 0−100% roots affected.

yPathogen recovery percentages represents the average percentage values of 4 plates cultured with 10 roots taken from each viburnum plant grown in pots. 

xMeans that do not share the same letter in a column are significantly different at P0.05, according to the Fisher’s protected least significant difference test.

Table 4. Colony forming units of Rhizoctonia solani grown on R. solani selective medium 

Treatment 

Biofumigation (BF) (4 weeks)zy

Biofumigation (BF) (2 weeks)zy

Before BF

24 hrs after BF 

4 weeks after BF 

24 weeks after BF

Before BF 

24 hrs after BF 

2 weeks after BF 

10 weeks after BF 

Amara mustard green        

3.66 ab

3.19 d

3.04 d

3.18 e

3.93 a

3.34 d

3.22 cd

3.27 d

Astro arugula

3.65 ab

3.44 bc

3.27 bc

3.36 de

3.90 a

3.43 d

3.26 cd

3.42 cd

Mighty mustard® pacific gold

3.68 ab

3.37 cd

3.18 cd

3.35 de

3.82 a

3.48 cd

3.19 d

3.40 d

Mustard

3.80 ab

3.43 bc

3.19 cd

3.42 cd

3.88 a

3.49 cd

3.38 c

3.42 cd

Oriental mustard

3.81 ab

3.54 b

3.46 b

3.56 bc

3.91 a

3.54 bcd

3.37 c

3.44 cd

Radish

3.65 ab

3.39 bcd

3.45 b

3.63 b

3.94 a

3.71 abc

3.59 b

3.59 bc

Dwarf essex rape

3.77 ab

3.55 b

3.47 b

3.57 bc

3.89 a

3.77 ab

3.63 b

3.69 b

Purple top forage turnips

3.62 b

3.31 cd

3.19 cd

3.34 de

3.91 a

3.48 cd

3.23 cd

3.42 cd

Non-inoculated control

2.00 c

2.00 e

2.10 e

2.20 f

2.30 b

2.30 e

2.42 e

2.75 e

Inoculated control

3.85 a

3.89 a

4.00 a

4.12 a

3.91 a

3.94 a

4.00 a

4.04 a

P-value 

<0.0001

<0.0001

<0.0001

<0.0001

<0.0001

<0.0001

<0.0001

<0.0001

zValues represent the log values of colony forming units (CFU) of fungi grown on R. solani selective medium. Colony counts were taken at 10-1 dilution series and colony forming units were calculated using CFU/ml = (no. of colonies x dilution factor) / volume of culture plate, formula and CFU values were converted into log values before analysis.

yMeans that do not share a letter are significantly different, according to the Fisher’s protected least significant difference test at P0.05.

Table 5. Height and width increase of viburnum (Viburnum odoratissimum) plants inoculated with Rhizoctonia solani during the experimental time period

Treatment

Biofumigation (4 weeks)zy

Biofumigation (2 weeks)zy

Height increase

Width  increase

Height  increase

Width   increase

Amara mustard green 

14.67 a

15.33 ab

7.00 ab

7.17 a

Astro arugula

14.33 a

21.67 a

8.00 a

5.83 a

Mighty mustard® pacific gold

7.67 bc

20.33 a

3.83 bcd

4.67 ab

Mustard

7.00 bc

12.33 bc

6.33 abc

4.75 ab

Oriental mustard

5.33 bc

8.67 bc

2.83 cd

4.67 ab

Radish

5.67 bc

11.00 bc

4.67 bcd

4.00 ab

Dwarf essex rape

6.67 bc

11.33 bc

7.33 ab

4.00 ab

Purple top forage turnips

8.67 abc

15.67 ab

4.00 bcd

6.33 a

Non-inoculated control 

10.00 ab

15.17 ab

6.33 abc

5.00 ab

Inoculated control

2.33 c

6.67 c

0.33 d

1.33 b

P-value 

0.0170

0.0090

0.0110

0.0180

zHeight and width measurements were taken in centimeter (cm).

 yMeans that do not share a letter are significantly different, according to the Fisher’s protected least significant difference test at P0.05.

Table 6. Whole plant fresh weight and root fresh weight of viburnum plants grown in Brassica cover crop incorporated soils with and without pathogen (Rhizoctonia solani) inoculation

Treatment

Biofumigation 

(4 weeks)zy

Biofumigation 

(2 weeks) zy

Whole plant fresh weight

Root fresh

weight

Whole plant fresh weight

Root fresh 

weight

Amara mustard green          

78.50 a

62.00 a

54.83 a

28.17 bcd

Astro arugula

72.00 ab

38.00 bcd

58.83 a

41.33 a

Mighty mustard® pacific gold

59.83 abc

47.17 ab

45.00 ab

25.00 bcd

Mustard

57.67 abc

36.17 bcd

52.33 ab

33.17 abc

Oriental mustard

46.33 bcd

29.17 bcd

39.83 bc

22.33 cde

Radish

43.33 cd

24.67 cd

36.33 bc

17.83 de

Dwarf essex rape

53.67 a-d

31.00 bcd

46.00 ab

35.33 ab

Purple top forage turnips

68.00 abc

42.00 bc

45.17 ab

23.33 cde

Non-inoculated control  

58.50 abc

40.17 bc

45.50 ab

29.00 bcd

Inoculated control

28.00 d

20.83 d

18.67 c

12.17 e

P-value 

0.0470

0.0110

0.0270

0.0020

zFresh weight measurements were taken in grams (g).

yMeans that do not share a letter are significantly different, according to the Fisher’s protected least significant difference test at P0.05.

Table 7. Phytophthora root rot disease severity on hydrangea (Hydrangea paniculata) and pathogen recovery from the roots grown in a soil biofumigated for 4 and 2 weeks 

Treatment

Biofumigation 

(4 weeks)

Biofumigation 

(2 weeks)

Disease severity 

(%)zx

Pathogen recovery from roots (%)yx

Disease severity 

(%)zx

Pathogen recovery from roots (%)yx

Amara mustard green          

23.42 c

28.33 c

23.42 d

25.83 bc

Astro arugula

15.08 cd

33.33 bc

23.42 d

29.17 bc

Mighty mustard® pacific gold

27.58 c

42.50 bc

31.75 bcd

35.00 ab

Mustard

23.42 c

40.00 bc

31.75 bcd

30.83 b

Oriental mustard

23.42 c

41.67 bc

40.08 bc

45.00 ab

Radish

56.75 ab

65.83 a

44.25 b

46.67 ab

Dwarf essex rape

15.08 cd

38.33 bc

27.58 cd

27.50 bc

Purple top forage turnips

48.42 b

46.67 b

35.92 bcd

32.50 b

Non-inoculated control  

8.67 d

0.00 d

2.25 e

6.67 c

Inoculated control

69.25 a

77.50 a

65.08 a

57.50 a

P-value

<0.0001

<0.0001

<0.0001

<0.0001

zRoot rot disease severity was recorded at the end of the trial for each plant using a scale of 0−100% roots affected.

yPathogen recovery percentages represents the average percentage values of 4 plates cultured with 10 roots taken from each hydrangea plant grown in pots. 

xMeans that do not share the same letter in a column are significantly different at P0.05, according to the Fisher’s protected least significant difference test.

Table 8. Colony forming units of Phytohthora nicotianae grown on PARPH-V8 medium

Treatment 

Biofumigation (BF) (4 weeks)zy

Biofumigation (BF) (2 weeks)zy

Before BF

24 hrs after BF

4 weeks after BF 

24 weeks after BF

Before BF

24 hrs after BF

2 weeks after BF

10 weeks after BF

Amara mustard green         

3.10 a

2.30 ef

2.16 de

2.30 f

3.21 a

2.59cd

2.30 de

2.10 cd

Astro arugula

3.11 a

2.49 de

2.10 e

2.30 f

3.20 a

2.46 de

2.16 ef

2.20 c

Mighty mustard® pacific gold

3.07 a

2.63 cde

2.46 cd

2.67 de

3.20 a

2.85 bc

2.69 c

2.93 b

Mustard

3.20 a

2.71 cd

2.20 de

2.59 def

3.17 a

2.20 e

2.10 ef

2.20 c

Oriental mustard

3.17 a

2.71 cd

2.54 c

2.77 cd

3.38 a

2.59 cd

2.52 cd

2.77 b

Radish

3.18 a

3.11 ab

3.19 ab

3.27 ab

3.31 a

3.26 a

3.27 ab

3.38 a

Dwarf essex rape

3.07 a

2.46 def

2.26 cde

2.43 ef

3.25 a

2.36 de

2.43 d

2.10 cd

Purple top forage turnips

3.19 a

2.90 bc

2.97 b

3.07 bc

3.26 a

3.16 ab

3.14 b

3.31 a

Non-inoculated control  

2.00 b

2.10 f

2.00 e

2.00 g

2.00 b

2.00 e

2.10 f

2.15 d

Inoculated control

3.15 a

3.28 a

3.33 a

3.39 a

3.33 a

3.39 a

3.42 a

3.45 a

P-value 

<0.0001

<0.0001

<0.0001

<0.0001

<0.0001

<0.0001

<0.0001

<0.0001

zValues represent the log values of colony forming units (CFU) of P. nicotianae grown on PARPH-V8 medium. Colony counts were taken at 10-1 dilution series and colony forming units were calculated using CFU/ml = (no. of colonies x dilution factor) / volume of culture plate, formula and CFU values were converted into log values before analysis.

yMeans that do not share a letter are significantly different, according to the Fisher’s protected least significant difference test at P0.05.

Table 9. Height and width increase of hydrangea (Hydrangea paniculata) plants inoculated with Phytophthora nicotianae during the experimental time period

Treatment

Biofumigation (4 weeks)zy

Biofumigation (2 weeks)zy

Height increase

Width   increase

Height  increase

Width   increase

Amara mustard green 

11.67 a 

17.67 a

8.33 a

14.17 a

Astro arugula

14.00 a

13.50 a

9.67 a

12.75 a

Mighty mustard® pacific gold

14.00 a

14.33 a

8.67 a

12.17 a

Mustard

12.00 a

15.67 a

8.00 a

11.50 a

Oriental mustard

13.00 a

10.50 a

5.67 a

10.83 a

Radish

10.00 a

10.33 a

7.67 a

9.17 a

Dwarf essex rape

16.00 a

16.83 a

8.00 a

10.83 a

Purple top forage turnips

10.33 a

11.50 a

8.33a

10.83 a

Non-inoculated control  

13.00 a

11.67 a

9.33 a

9.83 a

Inoculated control

7.00 a

8.83 a

2.67a

4.33 a 

P-value 

0.9300

0.6000

0.4600

0.3000

zHeight and width measurements were taken in centimeter (cm).

yMeans that do not share a letter are significantly different, according to the Fisher’s protected least significant difference test at P0.05.

Table 10. Whole plant fresh weight and root fresh weight of hydrangea plants grown in Brassica cover crop incorporated soils with and without pathogen (Phytophthora nicotianae) inoculation 

Treatment 

Biofumigation 

(4 weeks)zy

Biofumigation 

(2 weeks) zy

Whole plant fresh weight

Root fresh 

weight

Whole plant fresh weight

Root fresh

weight

Amara mustard green          

31.67 ab

22.17 ab

57.67 ab

50.67 a

Astro arugula

35.33 a

26.00 a

68.83 a

51.83 a

Mighty mustard® pacific gold

20.50 bcd

12.50 cd

44.67 ab

33.67 ab

Mustard

21.00 bcd

15.83 bcd

47.67 ab

34.50 ab

Oriental mustard

23.50 bcd

15.00 bcd

46.00 ab

31.50 ab

Radish

18.83 cd

10.00 cd

40.00 b

28.83 ab

Dwarf essex rape

34.33 a

24.00 ab

56.83 ab

39.33 a

Purple top forage turnips

20.33 bcd

15.50 bcd

48.33 ab

35.33 a

Non-inoculated control  

24.50 abc

17.83 abc

50.83 ab

36.33 a

Inoculated control

11.17 d

8.50 d

15.17 c

11.33 b

P-value 

0.017

0.008

0.025

0.086

yMeans that do not share a letter are significantly different, according to the Fisher’s protected least significant difference test at P0.05.

Table 11. Colony forming units of fluorescent Pseudomonads grown in Pseudomonads selective medium

Treatment

Biofumigation 

(4 weeks)zy

Biofumigation 

(2 weeks)zy

R. solani inoculated soil

P. nicotianae  inoculated soil

R. solani  inoculated soil

P. nicotianae  inoculated soil 

Amara mustard green

3.70 

3.88 

2.76 

3.27 

Astro arugula

3.73 

3.70 

3.69 

2.94 

Mighty mustard® pacific gold

3.64 

4.00 

3.30 

2.73 

Mustard

4.07 

4.13 

2.82 

2.97 

Oriental mustard

3.87 

3.98 

2.75 

3.21 

Radish

3.98 

4.10 

2.47 

2.74 

Dwarf essex rape

3.82 

3.95 

2.82 

2.85 

Purple top forage turnips

3.50 

4.08 

2.83 

3.03 

Non-inoculated control  

3.35 

3.16 

2.52 

2.32 

Inoculated control

3.98 

4.33

2.76 

2.27 

P-value

0.472

0.091

0.078

0.145

zValues represent the log values of colony forming units (CFU) of fluorescent Pseudomonads bacteria grown on Pseudomonads selective medium. Colony counts were taken at 10-1 dilution series and colony forming units were calculated using CFU/ml = (no. of colonies x dilution factor) / volume of culture plate, formula and CFU values were converted into log values before analysis.

yMeans that do not share a letter are significantly different, according to the Fisher’s protected least significant difference test at P0.05.

Table 12. Correlation between severity of diseases (%) caused by Rhizoctonia solani and Phytophthora nicotianae and fluorescent Pseudomonads log values in 4 weeks and 2 weeks biofumigation experiments 

 

Fluorescent Pseudomonads log values*

Biofumigation (4 weeks)

Biofumigation (2 weeks)

R. solani

P. nicotianae

R. solani

P. nicotianae

Disease severity 

R2=0.370

P= 0.044

R2=0.445

P= 0.014

R2= -0.141

P= 0.456

R2= -0.076

P= 0.691

*Negative values represent a negative correlation and values closer +1 or -1 indicates a strong relationship between growth data and root rot disease severity.

 
Participation Summary
1 Farmer participating in research

Educational & Outreach Activities

10 Consultations
1 Curricula, factsheets or educational tools
1 On-farm demonstrations
2 Published press articles, newsletters
2 Tours
5 Webinars / talks / presentations
1 Workshop field days

Participation Summary

60 Farmers
60 Ag professionals participated
Education/outreach description:

EDITOR REVIEWED ONLINE PUBLICATIONS

Baysal-Gurel, F., Simmons, T., Kabir, Md.N., Liyanapathiranage, P. 2017. Evaluation of biorational products and fungicides for the control of Phytophthora root rot of Hydrangea, 2016. Plant Disease Management Report No. 11:OT004. Online publication. The American Phytopathological Society, St. Paul, MN.

Baysal-Gurel, F., Simmons, T., Liyanapathiranage, P., Kabir, Md.N. 2017. Evaluation of biorational products and fungicides for the control of Rhizoctonia root rot of viburnum, 2016. Plant Disease Management Report No. 11:OT003. Online publication. The American Phytopathological Society, St. Paul, MN.

 

PUBLISHED ABSTRACTS

Kabir, Md N., Liyanapathiranage, P., Simmons, T., Baysal-Gurel, F. 2017. Evaluation of biorational products and fungicides for management of Phytophthora root rot of Hydrangea. TSU 39th Research Symposium. April 17-21, 2017. Nashville, Tenneessee.

Liyanapathiranage, P. and Baysal-Gurel, F. 2017. Pathogenicity of Rhizoctonia solani and Phytophthora nicotianae to Biofumigation Cover Crops (Brassica spp.). TSU 39th Research Symposium. April 17-21, 2017. Nashville, Tenneessee.

Liyanapathiranage, P., Kabir, Md N., Simmons, T., Baysal-Gurel, F. 2016. Evaluation of inoculation methods and inoculum level of Rhizoctonia solani to determine disease response. Tennessee Academy of Science- Annual Meeting. November 19, 2016. Austin Peay State University, Clarksville, Tennessee.

Liyanapathiranage, P., Kabir, Md N., Simmons, T., Baysal-Gurel, F.2017. Sustainable approaches for soilborne disease management in nursery production. The American Phytopathological Society. August 6-9, 2017. San Antonio, TX. 144-P. 

Dawadi, S., Addesso, K., O’Neal, P., Liyanapathiranage, P., Pandey, M., Baysal-Gurel, F.2017. Impact of cover crop usage on soilborne pathogens in a nursery production system. The American Phytopathological Society. August 6-9, 2017. San Antonio, TX. 149-P.

Liyanapathiranage, P., Kabir, Md N., Simmons, T., Baysal-Gurel, F. 2017. Pathogenicity of Rhizoctonia solani and Phytophthora nicotianae to Brassicaceae Biofumigant Cover Crops. The American Phytopathological Society. August 6-9, 2017. San Antonio, TX. 496-P. 

 

EXTENSION PUBLICATIONS

Baysal-Gurel, F., Kabir, Md N., and Blalock, A., 2016. Root diseases of hydrangeas. ANR-PATH-4-2016. TSU

 

ORAL PRESENTATIONS

Baysal-Gurel, F. 2016. Soil borne diseases of woody ornamentals in nursery production systems. Multistate Project S-1053- Ecological and Genetic Diversity of Soil-borne Pathogens and Indigenous Microflora Annual Meeting, McMinnville, TN October 19, 2016.

Liyanapathiranage, P. and Baysal-Gurel, F. 2016. Biofumigation for soil-borne disease management. Multistate Project S-1053- Ecological and Genetic Diversity of Soil-borne Pathogens and Indigenous Microflora Annual Meeting, McMinnville, TN October 19, 2016.

 

POSTER PRESENTATIONS

Baysal-Gurel, F.,and Liyanapathiranage, P. 2018. Sustainable Management of Soil-borne Diseases in Nursery Production. Our Farms, Our Future Conference. Apr 3-5, 2018. St. Louis, Missouri (Poster presentation- SSARE travel award).

 

FIELD DAY

Baysal-Gurel, F. 2017 Tennessee Nursery Field Day. TSU NRC McMinnville, TN. June 16, 2017.

Baysal-Gurel, F. 2018 Tennessee Nursery Field Day. TSU NRC McMinnville, TN. July 24, 2018.

 

THESIS

Liyanapathiranage, P. Sustainable management of soilborne diseases in nursery production. MS Thesis. Nashville, TN. 12/09/2017. (Major advisor: Fulya Baysal-Gurel)

Project Outcomes

2 Farmers reporting change in knowledge, attitudes, skills and/or awareness
2 Farmers changed or adopted a practice
1 Grant received that built upon this project
Project outcomes:

Soilborne diseases reduce plant performance; increase costs to the grower and cause potential ecological damage to the natural environment. Some soilborne pathogens may have broad host ranges and crop species may be susceptible to several different pathogens. The use of biofumigant cover crops has been explored most extensively in vegetable, fruit and flower production. However, their use and value have not been documented in woody ornamental nursery production. The objective of this study was to assess environmental friendly biofumigant cover crops for soilborne disease and improved plant growth. As the fisrt objective of the research inoculation methods and inoculum levels of Rhizoctonia solani was evaluated to determine disases response. Agar slurry inoculation method was showing a higher disease response when viburnum plants were inoculated. As all the tested three inoculum levels of the agar slurry method were showing similar root disease percentages, lowest inoculum level 1 petri dishes/L was selected and used for the following experiments. Fifteen biofumigant cover crops in the Brassicaceae family were evealuted for susceptibility to R. solani and Phytophthora nicotianae under greenhouse conditons. Oilseed radish, mustard, purple top forage turnip, astro arugula, mighty mustard® pacific gold, oriental mustard, dwarf essex rape Brassica, amara mustard green are showing low root rot disease percentages and diseases incidence in top soil which had pre-existing populations of R. solani and P. nicotinanae separately. These 8 cover crops were selected for the next experiment to evaluate the ability of them in releasing glucosinolate hydrolyze compoundsand control R. solani and P. nicotinanae.

Knowledge Gained:

With the introduction of the green revolution in 1960’s, traditional agriculture dramatically changed into modern agriculture with use of new technologies, increased chemical use, mechanization and specialization to maximize crop production to feed an increasing population. Even though these new implementations have had many positive effects in terms of production and efficiency, there have also been long-term negative effects on the environment and environmental sustainability along with other impacts to the social and economic conditions in rural farming communities. To address these problems, moving towards sustainable agriculture is becoming a trending solution. It provides solutions to major environmental and social issues while providing economically viable, innovative opportunities those who are into agriculture.

Nursery crops are produced on approximately 369,000 acres in the United States with Tennessee accounting for almost 10% of that production at 33,591 acres (USDA, 2014). Soil-borne diseases are a major limitation to the nursery production and conventionally used synthetic fungicides need to be applied every 7-14 days throughout the growing season to control outbreaks. But use of these fungicides disturbs the balance of the environment. Fungicides can cause adverse effects to human health, aquatic ecosystems and beneficial microorganisms in the soil. To control soil-borne diseases, while simultaneously maintaining the balance of the environment, some other environmentally friendly approaches like crop rotation, soil solarization, use of improved disease resistant varieties and biorational products can be used. Use of cover crops to biofumigate the soil is another approach that can be used because of its ability to release compounds with the ability to kill soil-borne pathogens. Biofumigant cover crop usage also increases the environmental sustainability compared to the synthetic fungicide usage. Once we apply synthetic fungicides to the soil it can take hours to years for the chemicals to completely degrade. Natural chemicals produced by the cover crops are degraded more quickly and can also be less harmful to the people who are handling these biofumigants. Again with fewer negative affects to handlers and the environment, use of biofumigants not only helps to protect crops now but also does in a way that conserves the environment for the future generation.

The use of biofumigant cover crops is a newer area of research in woody ornamental nursery production that has been previously explored most extensively in vegetable, fruit and flower production. This work will improve nursery production efficiency and reduce soil-borne disease pressure through applications of effective biofumigants in nursery production. Implementation of this project will contribute unique information on nursery production and innovative protocols by characterizing the impact of biofumigants on soil-borne plant pathogenic populations and demonstrating the role of microbial diversity and ecology in disease suppression. The information generated by these studies will provide sustainable management alternatives to growers combating soil-borne diseases.

Any opinions, findings, conclusions, or recommendations expressed in this publication are those of the author(s) and do not necessarily reflect the view of the U.S. Department of Agriculture or SARE.