This research was funded by the United States Department of Agriculture, Agricultural Research Service. Mention of trade names and commercial products in this article are solely for the purpose of providing specific information and do not imply recommendations or endorsements by the U.S. Department of Agriculture. USDA is an equal opportunity provider and employer. The authors have no conflict of interest to declare. Technical assistance was provided by Kristi Jordan.

1. Maintaining a Source of R. reniformis Inoculum
<ol>
	<li>Fill terra cotta clay pots (15 cm in diameter, 13.5 cm in height) with a steam pasteurized mix of 1-part sandy loam&#160;and 2-parts sand. Plant a susceptible tomato (Solanum lycopersicon) variety in each pot and place the pots in a glasshouse. 
	NOTE: Other susceptible plant varieties such as cotton can be used instead of tomato.</li>
	<li>Inoculate the tomato plants with vermiform reniform nematodes (see step 3.3). Maintain the plants in the glasshouse at a temperature of approximately 28 &#176;C.</li>
</ol>
2. Planting Cotton Genotypes for R. reniformis Resistance Evaluation
<ol>
	<li>Prepare soil by combining 2-parts fine sand with 1-part sandy loam&#160;collected from the field.</li>
	<li>Steam pasteurize the soil mixture to ensure that the soil is free of nematodes and soil-borne plant pathogens.</li>
	<li>Add the soil mixture to conical plastic pots (4 cm in diameter, 21 cm in height). Prior to filling the pots, place a ball of cotton in the bottom of the pot to prevent soil loss. Partially fill the pots with soil approximately 2 cm from the top.</li>
	<li>Prepare a plastic stake for each pot to designate the genotype to be planted.</li>
	<li>Plant seeds of the cotton genotypes selected for evaluation.
	<ol>
		<li>Plant a single seed in each pot for the evaluation of segregating populations, in which each seed represents a unique genotype.</li>
		<li>For cotton varieties or germplasm accessions, plant 2 to 3 seeds in a single pot to insure germination of at least one plant with the other seedlings removed from the pots prior to nematode inoculation. 
		Note: Alternatively, seeds can be germinated for 24 to 72 h prior to planting to minimize the number of pots with non-viable seeds.</li>
	</ol>
	</li>
	<li>Plant seeds of selected resistant and susceptible control genotypes. 
	Note: Control genotypes are replicated 5 to 10 times to assess natural variations inherent to the screening methodology.</li>
	<li>Fill pots with additional soil to cover the seed in each pot.</li>
	<li>Place the pots in the growth chamber. Maintain a constant temperature of 28 &#176;C, an ambient air temperature, for the growth chamber. Provide artificial lighting with a mixture of fluorescent and incandescent lamps with a 16 h photoperiod.</li>
	<li>Place water emitters in each pot, and water the pots twice per day using an automatic watering system. Adjust the watering system to supply additional water as the plant grows.</li>
</ol>
3. Nematode Inoculation of Cotton Plants and Preparation of Root Samples
<ol>
	<li>Extract vermiform reniform nematodes maintained on susceptible tomato plants (see step 1) using elutriation19 and centrifugal flotation20 methodologies the day before inoculation. Store the extracted nematodes at 4 &#176;C. 
	NOTE: Baermann funnel extraction21 is an alternate method for nematode extraction.</li>
	<li>Determine the number of nematodes extracted by counting the number of nematodes in a 100 &#181;L subsample and prepare a suspension of 1,000 nematodes/mL in tap water for inoculations.</li>
	<li>Inoculate the cotton seedlings 7 d after planting with the nematode suspension. Create a small depression in the soil next to the plant, and pipette 1 mL of the reniform nematode suspension into the depression.</li>
	<li>Remove plants from the pots 28 d after inoculation for nematode evaluation. 
	NOTE: At this stage, plants are approximately 15 cm tall with 4 to 6 fully expanded leaves.
	<ol>
		<li>Remove most of the fully expanded leaves from the plants using scissors prior to removing the plants from the pots.</li>
		<li>To remove plants from the pots, squeeze the pot and slide the soil out into the hand.</li>
		<li>Gently remove the soil from the roots by agitating the root system in tap water in a 10 L container. Briefly rinse the root system in a container of clean tap water.</li>
		<li>Remove the root system from the plant approximately 1 cm below the soil line using scissors.</li>
	</ol>
	</li>
	<li>Place the root system into a 120 mL plastic, non-sterile, disposable specimen container along with the plastic stake from the pot used for identification. 
	NOTE:&#160;Multiple samples are processed before proceeding to step 3.7. Proceed to step 4 for the vegetative propagation of the plant shoot.</li>
	<li>Prepare a 12.5% (v/v) solution of red food coloring17 in tap water to stain the nematodes attached to the root system.</li>
	<li>Add approximately 30 mL of the red food coloring solution to the root sample in the specimen container to completely cover the root system.</li>
	<li>Place the specimen container in a microwave oven and heat the root sample until the staining solution starts to boil. Remove the sample from the microwave oven and allow the sample to cool at room temperature.</li>
	<li>Decant the red food coloring solution from the root sample and add approximately 100 mL of tap water to the specimen container to remove excess stain. Place the cover on the specimen container and store the sample in a refrigerator at 4 &#176;C. Proceed to step 5 for evaluation of root infection. 
	NOTE:&#160;The protocol can be paused here before proceeding to step 5.</li>
</ol>
4. Vegetative Propagation to Recover Plants for Seed Production
<ol>
	<li>Place a ball of cotton in the bottom of a conical plastic pot (see step 2.3) and partially fill the pot with peat moss potting media. Then, place the vegetative shoot in the pot and firmly add potting media to fill the pot. Place a new labeled plastic stake in each pot to designate the cotton genotype.</li>
	<li>Place the tray of pots in a plastic container (73.6 cm length x 45.7 cm width x 15.2 cm height) with water and briefly water the plants to moisten the potting media. Place the pots in a growth chamber with a constant temperature of 28 &#176;C using a 16 h photoperiod. Add additional water to the plastic container as needed to maintain soil moisture.</li>
	<li>Transplant the plants to larger pots for seed production after approximately 30 d. Partially fill a 6 L plastic pot with potting media (see step 4.1), remove the plant from the small pot, place the plant into the 6 L pot, and firmly add potting media to fill the pot.</li>
	<li>Place the plants in a glasshouse and add water to moisten the potting media. Maintain the temperature in the glasshouse at approximately 28 &#176;C (artificial lighting is not required).
	<ol>
		<li>Water plants by hand as needed for approximately 30 d.</li>
		<li>When approximately 75% of the plants require daily watering, water plants daily using an automatic watering system. Adjust the automatic watering system to water more frequently as needed for plant growth.</li>
		<li>Add approximately 10 g of a slow-release fertilizer to each pot prior to the start of floral initiation.</li>
	</ol>
	</li>
	<li>Harvest plants at maturity and process the cotton seed samples to obtain seeds for further evaluation.
	<ol>
		<li>To harvest cotton seeds, remove the cotton from the open bolls on the plant by hand and place it in a labeled paper bag. The seeds are attached to the cotton fibers, which are removed in the following steps.</li>
		<li>Remove lint fibers from the seed samples using a 10-saw laboratory gin.</li>
		<li>Remove fuzz fibers from the seed samples using concentrated sulfuric acid. Neutralize seed samples in a 15% (v/v) solution of sodium carbonate, rinse the samples with tap water, and dry the samples in a forced air drier.</li>
		<li>Place the seed samples in labeled envelopes for storage.</li>
	</ol>
	</li>
</ol>
5. Evaluation of R. reniformis Root Infection
<ol>
	<li>Remove the root sample from the specimen container and count the number of female nematodes attached to the root system using a stereomicroscope (20X magnification). 
	NOTE:&#160;Only female reniform nematodes are capable of infecting plant roots.</li>
	<li>Place the root system on paper towels for approximately 10 min to remove excess moisture. Weigh the root system to determine the fresh root weight.</li>
	<li>Enter the nematode count and fresh root weight data into a computer spreadsheet program and calculate the number of females per gram of root.</li>
</ol>

<ol>
	<li>Heald, C. M., Robinson, A. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Survey+of+current+distribution+of+Rotylenchulus+reniformis.+in+the+United+States.">Survey of current distribution of Rotylenchulus reniformis. in the United States.</a> Journal of Nematology. 22 (4), 695-699 (1990).</li><li>Koenning, S. R., Wrather, J. A., Kirkpatrick, T. L., Walker, N. R., Starr, J. L., Mueller, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plant-parasitic+nematodes+attacking+cotton+in+the+United+States:+old+and+emerging+production+challenges.">Plant-parasitic nematodes attacking cotton in the United States: old and emerging production challenges.</a> Plant Disease. 88 (2), 100-113 (2004).</li><li>Robinson, A. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reniform+in+U.S.+cotton:+when,+where,+why,+and+some+remedies.">Reniform in U.S. cotton: when, where, why, and some remedies.</a> Annual Review of Phytopathology. 45, 263-288 (2007).</li><li>Robinson, A. F., Inserra, R. N., Caswell-Chen, E. P., Vovlas, N., Troccoli, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rotylenchulus+species:+identification,+distribution,+host+ranges,+and+crop+plant+resistance.">Rotylenchulus species: identification, distribution, host ranges, and crop plant resistance.</a> Nematropica. 27 (2), 127-180 (1997).</li><li>Davis, R. F., Koenning, S. R., Kemerait, R. C., Cummings, T. D., Hurley, W. 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E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+sources+of+resistance+to+the+reniform+(Rotylenchulus+reniformis)+and+root-knot+(Meloidogyne+incognita)+nematode+in+upland+(Gossypium+hirsutum+L.)+and+sea+island+(G.+barbadense+L.)+cotton.">New sources of resistance to the reniform (Rotylenchulus reniformis) and root-knot (Meloidogyne incognita) nematode in upland (Gossypium hirsutum L.) and sea island (G. barbadense L.) cotton.</a> Journal of Cotton Science. 8 (3), 191-197 (2004).</li><li>Robinson, A. F., Percival, A. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Resistance+to+Meloidogyne+incognita+race+3+and+Rotylenchulus+reniformis+in+wild+accessions+of+Gossypium+hirsutum+and+G.+barbadense+from+Mexico.">Resistance to Meloidogyne incognita race 3 and Rotylenchulus reniformis in wild accessions of Gossypium hirsutum and G. barbadense from Mexico.</a> Journal of Nematology. 29 (4), 746-755 (1997).</li><li>Stetina, S. R., Young, L. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparisons+of+female+and+egg+assays+to+identify+Rotylenchulus+reniformis+resistance+in+cotton.">Comparisons of female and egg assays to identify Rotylenchulus reniformis resistance in cotton.</a> Journal of Nematology. 38 (3), 326-332 (2006).</li><li>Usery, S. R., Lawrence, K. S., Lawrence, G. W., Burmester, C. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+cotton+cultivars+for+resistance+and+tolerance+to+Rotylenchulus+reniformis.">Evaluation of cotton cultivars for resistance and tolerance to Rotylenchulus reniformis.</a> Nematropica. 35 (2), 121-133 (2005).</li><li>Yik, C. -. P., Birchfield, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Resistant+germplasm+in+Gossypium+species+and+related+plants+to+Rotylenchulus+reniformis.">Resistant germplasm in Gossypium species and related plants to Rotylenchulus reniformis.</a> Journal of Nematology. 16 (2), 146-153 (1984).</li><li>Stetina, S. R., Erpelding, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gossypium+arboreum+accessions+resistant+to+Rotylenchulus+reniformis.">Gossypium arboreum accessions resistant to Rotylenchulus reniformis.</a> Journal of Nematology. 48 (4), 223-230 (2016).</li><li>Thies, J. A., Merrill, S. B., Corley, E. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Red+food+coloring+stain:+new,+safer+procedures+for+staining+nematodes+in+roots+and+egg+masses+on+root+surfaces.">Red food coloring stain: new, safer procedures for staining nematodes in roots and egg masses on root surfaces.</a> Journal of Nematology. 34 (2), 179-181 (2002).</li><li>Erpelding, J. E., Stetina, S. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+characterization+of+reniform+nematode+resistance+for+Gossypium+arboreum+accession+PI+417895.">Genetic characterization of reniform nematode resistance for Gossypium arboreum accession PI 417895.</a> Plant Breeding. 137 (1), 81-88 (2018).</li><li>Byrd, D. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two+semi-automatic+elutriators+for+extracting+nematodes+and+certain+fungi+from+soil.">Two semi-automatic elutriators for extracting nematodes and certain fungi from soil.</a> Journal of Nematology. 8 (3), 206-212 (1976).</li><li>Jenkins, W. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+rapid+centrifugal-flotation+technique+for+separating+nematodes+from+soil.">A rapid centrifugal-flotation technique for separating nematodes from soil.</a> Plant Disease Reporter. 48 (9), 692 (1964).</li><li>Robinson, A. F., Heald, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Carbon+dioxide+and+temperature+gradients+in+Baermann+funnel+extraction+of+Rotylenchulus+reniformis.">Carbon dioxide and temperature gradients in Baermann funnel extraction of Rotylenchulus reniformis.</a> Journal of Nematology. 23 (1), 28-38 (1991).</li><li>Williams, C., Gilman, D. F., Fontenot, D. 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The authors have nothing to disclose.

An effective screening protocol is required for 1) the identification of R. reniformis resistant cotton genotypes in order to evaluate the genetics of resistance and 2) the breeding of resistant varieties. Most protocols assess R. reniformis population densities or reproduction rates by extracting vermiform nematodes or eggs from the cotton root system or potting soil8,11,12,<sup cl...

Rotylenchulus reniformis (Linford and Oliveira), commonly referred to as the reniform nematode, is one of the major parasitic nematode species present in soils of the southeastern United States1,2,3. The nematode is an obligate, sedentary semi-endoparasite requiring a host plant to complete its life cycle2,4. Vermiform preadult female nematodes penetrate the host root system to establish a feeding site in the stele2,3. As the nematode feeds and matures, the posterior portion remaining outside of the host root will swell upon egg production, forming a characteristic kidney shape (Figure 1). Rotylenchulus reniformis is capable of feeding on the root system of more than 300 plant species, including cotton4. Upland cotton (Gossypium hirsutum L.) is widely cultivated in the southeastern United States, but the lack of R. reniformis&#160;resistant varieties hinders nematode management2,3. Management strategies such as nematicide treatment and rotation with non-host crop species have been used to reduce soil R. reniformis population densities5,6, but seed cotton yield losses can commonly range from 1 to 5%2. Symptoms of R. reniformis infection can include plant stunting, suppressed root growth, nutritional deficiencies, fruit abortion, and delayed maturity2. However, symptoms may not be apparent due to the uniformity of symptoms across the field; therefore, approaches to assess R. reniformis infection are needed to identify and develop resistant upland cotton varieties. Evaluation of R. reniformis resistance in cotton is considered difficult7, because the infected root system may appear normal even though the plant may show symptoms of infection8.
An effective nematode screening protocol is required for the identification of R. reniformis&#160;resistant accessions from the cotton germplasm collection, and for the determination of the resistance genetics for these accessions. Such a protocol will aid in the transfer of resistance genes to upland cotton. Various bioassay methods have been used to assess R. reniformis infection in cotton8,9,10,11,12,13,14,15. In general, two major approaches have been used for the identification of R. reniformis resistant cotton genotypes. The most frequently used approach involves extracting eggs and/or vermiform nematodes from infected plants or soil8,11,12,14,15. The general methodology for this approach involves planting seeds for the individual cotton genotypes in separate pots, allowing the seedlings to develop for 7 to 14 days, inoculating the seedlings by adding a mixture of vermiform stages of R. reniformis to the soil, and allowing the nematodes to infect the root system for 30 to 60 days. Next, vermiform nematodes and/or eggs are extracted from the infected root system of each plant or from the potting soil. The number of extracted nematodes or eggs is then determined to estimate the population density and reproduction rate, which are compared to control genotypes in order to identify resistant genotypes.
An alternative approach, as described here, involves microscopically examining the cotton root system that has been infected with nematodes to determine the number of female nematodes parasitizing the roots10,16. Similar to other approaches, cotton genotypes are planted in separate pots and inoculated with vermiform nematodes approximately 7 days after planting. Within 30 days after inoculation, the root system is removed from individual plants and the soil is rinsed from the roots. Next, the nematodes attached to the root system are stained with red food coloring17, and roots are microscopically examined to determine the number of infection sites with resistant cotton genotypes (identified based on the number of nematodes per gram of root) compared to a susceptible control16. This second approach has the advantage of increased throughput by reducing the number of days required for evaluation and increasing the number of individual genotypes evaluated in a single experiment. Screening methodologies that evaluate population density or reproduction rate are often more time-consuming than those based on visual observations of infection signs7. However, one limitation of this approach is that host-plant resistance that hinders nematode reproduction as determined by egg production is not assessed13.
Screening protocols for R. reniformis resistance often destroy the root system during evaluation7 and involve the vegetative shoot being discarded. To overcome this limitation, a method of vegetative propagation has been developed to allow the recovery of plants for seed production18. After removal of the root system for nematode evaluation, the vegetative shoot is planted in potting soil to allow the root system to regrow. This method has broad applications for most R. reniformis screening protocols. A simple and rapid method of vegetative propagation is of critical importance for breeding R. reniformis resistant upland cotton varieties, where the recovery of the progeny is required to advance resistant genotypes to the next generation.
A protocol is presented for the large-scale screening of cotton genotypes for reniform nematode resistance. The goal is to develop a simple and rapid non-destructive screening method to evaluate cotton breeding populations for nematode resistance in order to aid in the breeding of resistant upland cotton varieties. Using this protocol, data are typically obtained within 35 days, with more than 300 genotypes evaluated in a single experiment. Data are presented for resistant and susceptible genotypes to illustrate the variation commonly observed using these methods.

<table><tbody><tr><td>Ray Leach Cone-tainer</td><td>Stuewe and Sons Inc.</td><td>SC10U</td><td></td></tr><tr><td>Cone-tainer tray</td><td>Stuewe and Sons Inc.</td><td>RL98</td><td></td></tr><tr><td>Sand</td><td>various</td><td></td><td></td></tr><tr><td>Cotton balls</td><td>various</td><td></td><td></td></tr><tr><td>Pylon 4 inch plant labels (4 in L x 5/8 in W)</td><td>Pylon Platics</td><td>L-4-W</td><td>Any brand or vendor is acceptible.</td></tr><tr><td>4 oz. specimen containers</td><td>Fisher Scientific</td><td>16-320-731</td><td>Any brand or vendor is acceptible.</td></tr><tr><td>Red food coloring</td><td>McCormick &amp;amp; Co., Inc.</td><td></td><td></td></tr><tr><td>1 mL Pipet tips</td><td>various</td><td></td><td></td></tr><tr><td>10 L container</td><td>various</td><td></td><td>Inexpensive buckets work well.</td></tr><tr><td>6 L pots</td><td>Nursery Supplies Inc.</td><td>Poly-Tainer-Can No2A</td><td>Any brand or vendor is acceptible. Different size pots can be used</td></tr><tr><td>Potting media</td><td>Sun Gro Horticulture</td><td>Metro-Mix 360</td><td>Any brand or vendor is acceptible.</td></tr><tr><td>Fertilizer</td><td>Everris NA Inc.</td><td>Osmocote Plus</td><td>Any brand or vendor is acceptible.</td></tr><tr><td>Plastic container (73.6 cm L x 45.7 cm W x 15.2 cm D)</td><td>Rubbermaid</td><td>3O29&amp;nbsp;</td><td>Any brand or vendor is acceptible.</td></tr></tbody></table>

screening cotton genotypes for reniform nematode resistance

A rapid non-destructive reniform nematode (Rotylenchulus reniformis) screening protocol is needed for the development of resistant cotton (Gossypium hirsutum) varieties to improve nematode management. Most protocols involve extracting vermiform nematodes or eggs from the cotton root system or potting soil to determine population density or reproduction rate. These approaches are generally time-consuming with a small number of genotypes evaluated. An alternative approach is described here in which the root system is visually examined for nematode infection. The protocol involves inoculating cotton seedling 7 days after planting with vermiform nematodes and determining the number of females attached to the root system 28 days after inoculation. Data are expressed as the number of females per gram of fresh root weight to adjust for variation in root growth. The protocol provides an excellent method for evaluating host-plant resistance associated with the ability of the nematode to establish an infection site; however, resistance that hinders nematode reproduction is not assessed. As with other screening protocols, variation is commonly observed in nematode infection among individual genotypes within and between experiments. Data are presented to illustrate the range of variation observed using the protocol. To adjust for this variation, control genotypes are included in experiments. Nonetheless, the protocol provides a simple and rapid method to evaluate host-plant resistance. The protocol has been successfully used to identify resistant accessions from the G. arboreum germplasm collection and evaluate segregating populations of more than 300 individuals to determine the genetics of resistance. A vegetative propagation method for recovering plants for resistance breeding was also developed. After removal of the root system for nematode evaluation, the vegetative shoot is replanted to allow the development of a new root system. More than 95% of the shoots typically develop a new root system with plants reaching maturity.

Rotylenchulus reniformis infection of the root system for two varieties is presented in Figure 1. Relatively fewer female reniform nematodes are able to establish a feeding site for the resistant cotton genotype compared to the susceptible genotype. Variation in root growth is common between accessions, as illustrated in Figure 2. This variation as measured by fresh root weight can also be observed between plants of the ...

Watch this Scientific Journal Video about Screening Cotton Genotypes for Reniform Nematode Resistance at JoVE.com

Screening Cotton Genotypes for Reniform Nematode Resistance

Here, a protocol is presented for the rapid non-destructive screening of cotton genotypes for reniform nematode resistance. The protocol involves visually examining the roots of nematode-infected cotton seedlings to determine infection response. The vegetative shoot from each plant is then propagated to recover plants for seed production.

A rapid non-destructive reniform nematode (Rotylenchulus reniformis) screening protocol is needed for the development of resistant cotton (Gossypium ...

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JoVE Journal

Genetics

10.3K Views.  United States Department of Agriculture. Deze methode kan helpen bij de ontwikkeling van reniforme nematodenbestendige katoenrassen. Door de identificatie van resistente genotypen van andere katoensoorten en de evaluatie van fokpopulaties. Het belangrijkste voordeel van deze techniek is dat het een eenvoudige en niet-destructieve methode biedt voor reniform nematode screening.Begin met het plaatsen van een bal van katoen in de bodem van een vier centimeter diameter, 21 centrum hoogte kegelvormige plastic pot per zaad. En het ...

Video: Screening Cotton Genotypes for Reniform Nematode Resistance

A Rapid and Efficient Method for Assessing Pathogenicity of Ustilago maydis on Maize and Teosinte Lines

Maize is a major cereal crop worldwide. However, susceptibility to biotrophic pathogens is the primary constraint to increasing productivity. U. maydis is a biotrophic fungal pathogen and the causal agent of corn smut on maize. This disease is responsible for significant yield losses of approximately $1.0 billion annually in the U.S.1 Several methods including crop rotation, fungicide application and seed treatments are currently used to control corn smut2. However, host resistance is the only practical method for managing corn smut. Identification of crop plants including maize, wheat, and rice that are resistant to various biotrophic pathogens has significantly decreased yield losses annually3-5. Therefore, the use of a pathogen inoculation method that efficiently and reproducibly delivers the pathogen in between the plant leaves, would facilitate the rapid identification of maize lines that are resistant to U. maydis. As, a first step toward indentifying maize lines that are resistant to U. maydis, a needle injection inoculation method and a resistance reaction screening method was utilized to inoculate maize, teosinte, and maize x teosinte introgression lines with a U. maydis strain and to select resistant plants.
Maize, teosinte and maize x teosinte introgression lines, consisting of about 700 plants, were planted, inoculated with a strain of U. maydis, and screened for resistance. The inoculation and screening methods successfully identified three teosinte lines resistant to U. maydis. Here a detailed needle injection inoculation and resistance reaction screening protocol for maize, teosinte, and maize x teosinte introgression lines is presented. This study demonstrates that needle injection inoculation is an invaluable tool in agriculture that can efficiently deliver U. maydis in between the plant leaves and has provided plant lines that are resistant to U. maydis that can now be combined and tested in breeding programs for improved disease resistance.

A Rapid and Efficient Method for Assessing Pathogenicity of Ustilago maydis on Maize and Teosinte Lines

The use of a needle injection method to inoculate maize and teosinte plants with the biotrophic pathogen Ustilago maydis is described. The needle injection inoculation method facilitates the controlled delivery of the fungal pathogen in between the plant leaves where the pathogen enters the plant through the formation of appresoria. This method is highly efficient, enabling reproducible inoculations with U. maydis.

Maize is a major cereal crop worldwide. However, susceptibility to biotrophic pathogens is the primary constraint to increasing productivity. U. maydis is ...

Environment

Semi-High Throughput Screening for Potential Drought-tolerance in Lettuce (Lactuca sativa) Germplasm Collections

This protocol describes a method by which a large collection of the leafy green vegetable lettuce (Lactuca sativa L.) germplasm was screened for likely drought-tolerance traits. Fresh water availability for agricultural use is a growing concern across the United States as well as many regions of the world. Short-term drought events along with regulatory intervention in the regulation of water availability coupled with the looming threat of long-term climate shifts that may lead to reduced precipitation in many important agricultural regions has increased the need to hasten the development of crops adapted for improved water use efficiency in order to maintain or expand production in the coming years. This protocol is not meant as a step-by-step guide to identifying at either the physiological or molecular level drought-tolerance traits in lettuce, but rather is a method developed and refined through the screening of thousands of different lettuce varieties. The nature of this screen is based in part on the streamlined measurements focusing on only three water-stress indicators: leaf relative water content, wilt, and differential plant growth following drought-stress. The purpose of rapidly screening a large germplasm collection is to narrow the candidate pool to a point in which more intensive physiological, molecular, and genetic methods can be applied to identify specific drought-tolerant traits in either the lab or field. Candidates can also be directly incorporated into breeding programs as a source of drought-tolerance traits.

Semi-High Throughput Screening for Potential Drought-tolerance in Lettuce Lactuca sativa Germplasm Collections

This protocol was developed to screen a large germplasm collection of the leafy vegetable lettuce (Lactuca sativa L.) for drought-tolerance, in order to identify a small candidate pool of lettuce for use in physiological, molecular, and genetic studies to identify underlying drought-tolerance traits along with breeding programs.

This protocol describes a method by which a large collection of the leafy green vegetable lettuce (Lactuca sativa L.) germplasm was screened for likely ...

Protocols for Robust Herbicide Resistance Testing in Different Weed Species

Robust protocols to test putative herbicide resistant weed populations at whole plant level are essential to confirm the resistance status. The presented protocols, based on whole-plant bioassays performed in a greenhouse, can be readily adapted to a wide range of weed species and herbicides through appropriate variants. Seed samples from plants that survived a field herbicide treatment are collected and stored dry at low temperature until used. Germination methods differ according to weed species and seed dormancy type. Seedlings at similar growth stage are transplanted and maintained in the greenhouse under appropriate conditions until plants have reached the right growth stage for herbicide treatment. Accuracy is required to prepare the herbicide solution to avoid unverifiable mistakes. Other critical steps such as the application volume and spray speed are also evaluated. The advantages of this protocol, compared to others based on whole plant bioassays using one herbicide dose, are related to the higher reliability and the possibility of inferring the resistance level. Quicker and less expensive in vivo or in vitro diagnostic screening tests have been proposed (Petri dish bioassays, spectrophotometric tests), but they provide only qualitative information and their widespread use is hindered by the laborious set-up that some species may require. For routine resistance testing, the proposed whole plant bioassay can be applied at only one herbicide dose, so reducing the costs.

A robust and flexible approach to confirm herbicide resistance in weed populations is presented. This protocol allows the herbicide resistance levels to be inferred and applied to a wide range of weed species and herbicides with minor adaptations.

Robust protocols to test putative herbicide resistant weed populations at whole plant level are essential to confirm the resistance status. The presented ...

High and Low Throughput Screens with Root-knot Nematodes Meloidogyne spp.

Root-knot nematodes (genus Meloidogyne) are obligate plant parasites. They are extremely polyphagous and considered one of the most economically important plant parasitic nematodes. The microscopic second-stage juvenile (J2), molted once in the egg, is the infective stage. The J2s hatch from the eggs, move freely in the soil within a film of water, and locate root tips of suitable plant species. After penetrating the plant root, they migrate towards the vascular cylinder where they establish a feeding site and initiate feeding using their stylets. The multicellular feeding site is comprised of several enlarged multinuclear cells called 'giant cells' which are formed from cells that underwent karyokinesis (repeated mitosis) without cytokinesis. Neighboring pericycle cells divide and enlarge in size giving rise to a typical gall or root knot, the characteristic symptom of root-knot nematode infection. Once feeding is initiated, J2s become sedentary and undergo three additional molts to become adults. The adult female lays 150-250 eggs in a gelatinous matrix on or below the surface of the root. From the eggs new infective J2s hatch and start a new cycle. The root-knot nematode life cycle is completed in 4-6 weeks at 26-28&deg;C.
Here we present the traditional protocol to infect plants, grown in pots, with root-knot nematodes and two methods for high-throughput assays. The first high-throughput method is used for plants with small seeds such as tomato while the second is for plants with large seeds such as cowpea and common bean. Large seeds support extended seedling growth with minimal nutrient supplement. The first high throughput assay utilizes seedlings grown in sand in trays while in the second assay plants are grown in pouches in the absence of soil. The seedling growth pouch is made of a 15.5 x 12.5cm paper wick, folded at the top to form a 2-cm-deep trough in which the seed or seedling is placed. The paper wick is contained inside a transparent plastic pouch. These growth pouches allow direct observation of nematode infection symptoms, galling of roots and egg mass production, under the surface of a transparent pouch. Both methods allow the use of the screened plants, after phenotyping, for crossing or seed production. An additional advantage of the use of growth pouches is the small space requirement because pouches are stored in plastic hanging folders arranged in racks.

High and Low Throughput Screens with Root-knot Nematodes Meloidogyne spp.

Two distinct methods to screen plants with root-knot nematodes are described. The described approaches include high-throughput screens with nematodes in a nondestructive manner facilitating the use of these plants in breeding programs.

Root-knot nematodes (genus Meloidogyne) are obligate plant parasites. They are extremely polyphagous and considered one of the most economically important ...

Immunology and Infection

VIGS-Mediated Forward Genetics Screening for Identification of Genes Involved in Nonhost Resistance

Nonhost disease resistance of plants against bacterial pathogens is controlled by complex defense pathways. Understanding this mechanism is important for developing durable disease-resistant plants against wide range of pathogens. Virus-induced gene silencing (VIGS)-based forward genetics screening is a useful approach for identification of plant defense genes imparting nonhost resistance. Tobacco rattle virus (TRV)-based VIGS vector is the most efficient VIGS vector to date and has been efficiently used to silence endogenous target genes in Nicotiana benthamiana.

	In this manuscript, we demonstrate a forward genetics screening approach for silencing of individual clones from a cDNA library in N. benthamiana and assessing the response of gene silenced plants for compromised nonhost resistance against nonhost pathogens, Pseudomonas syringae pv. tomato T1, P. syringae pv. glycinea, and X. campestris pv. vesicatoria. These bacterial pathogens are engineered to express GFPuv protein and their green fluorescing colonies can be seen by naked eye under UV light in the nonhost pathogen inoculated plants if the silenced target gene is involved in imparting nonhost resistance. This facilitates reliable and faster identification of gene silenced plants susceptible to nonhost pathogens. Further, promising candidate gene information can be known by sequencing the plant gene insert in TRV vector. Here we demonstrate the high throughput capability of VIGS-mediated forward genetics to identify genes involved in nonhost resistance. Approximately, 100 cDNAs can be individually silenced in about two to three weeks and their relevance in nonhost resistance against several nonhost bacterial pathogens can be studied in a week thereafter. In this manuscript, we enumerate the detailed steps involved in this screening. VIGS-mediated forward genetics screening approach can be extended not only to identifying genes involved in nonhost resistance but also to studying genes imparting several biotic and abiotic stress tolerances in various plant species.

Virus-induced gene silencing is an useful tool for identifying genes involved in nonhost resistance of plants. We demonstrate the use of bacterial pathogens expressing GFPuv in identifying gene silenced plants susceptible to nonhost pathogens. This approach is easy, fast and facilitates large scale screening and similar protocol can be applied to studying various other plant-microbe interactions.


	Nonhost disease resistance of plants against bacterial pathogens is controlled by complex defense pathways. Understanding this mechanism is important ...

RNAi Screening to Identify Postembryonic Phenotypes in C. elegans

C. elegans has proven to be a valuable model system for the discovery and functional characterization of many genes and gene pathways1. More sophisticated tools and resources for studies in this system are facilitating continued discovery of genes with more subtle phenotypes or roles. 
Here we present a generalized protocol we adapted for identifying C. elegans genes with postembryonic phenotypes of interest using RNAi2. This procedure is easily modified to assay the phenotype of choice, whether by light or fluorescence optics on a dissecting or compound microscope. This screening protocol capitalizes on the physical assets of the organism and molecular tools the C. elegans research community has produced. As an example, we demonstrate the use of an integrated transgene that expresses a fluorescent product in an RNAi screen to identify genes required for the normal localization of this product in late stage larvae and adults. First, we used a commercially available genomic RNAi library with full-length cDNA inserts. This library facilitates the rapid identification of multiple candidates by RNAi reduction of the candidate gene product. Second, we generated an integrated transgene that expresses our fluorecently tagged protein of interest in an RNAi-sensitive background. Third, by exposing hatched animals to RNAi, this screen permits identification of gene products that have a vital embryonic role that would otherwise mask a post-embryonic role in regulating the protein of interest. Lastly, this screen uses a compound microscope equipped for single cell resolution.

RNAi Screening to Identify Postembryonic Phenotypes in C. elegans

We describe a sensitized method to identify postembryonic regulators of protein expression and localization in C. elegans using an RNAi-based genomic screen and an integrated transgene that expresses a functional, fluorescently tagged protein.

C. elegans has proven to be a valuable model system for the discovery and functional characterization of many genes and gene pathways1. More sophisticated ...

Biology

Screening of Tobacco Genotypes for Phytophthora nicotianae Resistance

Black shank, caused by the oomycetes Phytophthora nicotianae, is destructive to tobacco, and this pathogen is highly pathogenic to many solanaceous crops. P. nicotianae is well adapted to high temperatures; therefore, research on this pathogen is gaining importance in agriculture worldwide because of global warming. P. nicotianae-resistant varieties of tobacco plants are commonly screened by inoculation with oat grains colonized by P. nicotianae and monitoring for the disease symptoms. However, it is difficult to quantify the inoculation intensity since accurate inoculation is crucial in this case. This study aimed to develop an efficient and reliable method for evaluating the resistance of tobacco to infection with P. nicotianae. This method has been successfully used to identify resistant varieties, and the inoculation efficiency was confirmed by real-time PCR. The resistance evaluation method presented in this study is efficient and practical for precision breeding, as well as molecular mechanism research.

Screening of Tobacco Genotypes for Phytophthora nicotianae Resistance

Here, a protocol is presented for the efficient and accurate screening of tobacco genotypes for Phytophthora nicotianae resistance in seedlings. This is a practical approach for precision breeding, as well as molecular mechanism research.

Black shank, caused by the oomycetes Phytophthora nicotianae, is destructive to tobacco, and this pathogen is highly pathogenic to many solanaceous crops. ...

A Contrast of Three Inoculation Techniques used to Determine the Race of Unknown Fusarium oxysporum f.sp. niveum Isolates

Fusarium wilt of watermelon (Citrullus lanatus), caused by Fusarium oxysporum f. sp. niveum (Fon), has reemerged as a major production constraint in the southeastern USA, especially in Florida. Deployment of integrated pest management strategies, such as race-specific resistant cultivars, requires information on the diversity and population density of the pathogen in growers' fields. Despite some progress in developing molecular diagnostic tools to identify pathogen isolates, race determination often requires bioassay approaches.
Race typing was conducted by root-dip inoculation, infested kernel seeding method, and the modified tray-dip method with each of the four watermelon differentials (Black Diamond, Charleston Grey, Calhoun Grey, Plant Introduction 296341-FR). Isolates are assigned a race designation by calculation of disease incidence five weeks after inoculation. If less than 33% of the plants for a particular cultivar were symptomatic, they were categorized as resistant. Those cultivars with incidence greater than 33% were regarded as susceptible. This paper describes three different methods of inoculation to ascertain race, root-dip, infested kernel, and modified tray-dip inoculation, whose applications vary according to the experimental design.

A Contrast of Three Inoculation Techniques used to Determine the Race of Unknown Fusarium oxysporum f.sp. niveum Isolates

Managing Fusarium wilt of watermelon requires knowledge of the pathogen races present. Here, we describe the root-dip, infested kernel seeding, and modified tray-dip inoculation methods to demonstrate their efficacy in race-typing of the pathogenic fungus Fusarium oxysporum f. sp niveum (Fon).

Fusarium wilt of watermelon (Citrullus lanatus), caused by Fusarium oxysporum f. sp. niveum (Fon), has reemerged as a major production constraint in the ...

Culturing and Screening the Plant Parasitic Nematode Ditylenchus dipsaci

Plant-parasitic nematodes (PPNs) destroy over 12% of global food crops every year, which equates to roughly 157 billion dollars (USD) lost annually. With a growing global population and limited arable land, controlling PPN infestation is critical for food production. Compounding the challenge of maximizing crop yields are the mounting restrictions on effective pesticides because of a lack of nematode selectivity. Hence, developing new and safe chemical nematicides is vital to food security. In this protocol, the culture and collection of the PPN species Ditylenchus dipsaci are demonstrated. D. dipsaci is both economically damaging and relatively resistant to most modern nematicides. The current work also explains how to use these nematodes in screens for novel small molecule nematicides and reports on data collection and analysis methodologies. The demonstrated pipeline affords a throughput of thousands of compounds per week and can be easily adapted for use with other PPN species such as Pratylenchus penetrans. The techniques described herein can be used to discover new nematicides, which may, in turn, be further developed into highly selective commercial products that safely combat PPNs to help feed an increasingly hungry world.

Culturing and Screening the Plant Parasitic Nematode Ditylenchus dipsaci

The present protocol describes a reliable and straightforward method for culturing, collecting, and screening Ditylenchus dipsaci.

Plant-parasitic nematodes (PPNs) destroy over 12% of global food crops every year, which equates to roughly 157 billion dollars (USD) lost annually. With ...

Genetic Screens

Genetic screens are critical tools for defining gene function and understanding gene interactions. Screens typically involve mutating genes and then assessing the affected organisms for phenotypes of interest. The process can be &#8220;forward&#8221;, where mutations are generated randomly to identify unknown genes responsible for the phenotypes, or it can be &#8220;reverse&#8221;, where specific genes are targeted for mutation to observe what phenotypes are produced.Here, JoVE reviews various types of genetic screens, including those that depend on either loss-of-function or gain-of-function mutations, which respectively decrease or increase the activity of genes. We then explore general protocols for forward and reverse screens in a popular model organism, the nematode worm. Finally, we highlight how screens are applied in research today, for example to better understand gene interactions that may contribute to neurodegenerative diseases.

Forward and Reverse Genetic Screens

Genetic screens are critical tools for defining gene function and understanding gene interactions. Screens typically involve mutating genes and then ...

Screening Cotton Genotypes for Reniform Nematode Resistance

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