Research in Kaloshian lab is funded by a grant from United States National Institute of Food and Agriculture (2007-35607-17765). Research in Roberts lab is funded by grants from United States Agency for International Development (GDG-G-00-02-00012-00 and EDH-A-00-07-00005) and the California Dry Bean Advisory Board.

1. Tomato Seedling Growth in Pots and Trays
<ol>
 <li>For pot assays, plant tomato seeds in a single pot in an organic-rich soil such as Sunshine mix. Maintain in a greenhouse at 22-28&deg;C. After germination, fertilize once a week with Miracle-Gro.</li>
 <li>About two weeks after germination, at the two true-leaf stage, transplant individually the seedlings into pots (10 cm diameter and 17 cm deep) filled with sterile sandy soil containing 90% sand and 10% organic mix. Add Osmocote slow release fertilizer and maintain in a greenhouse at 22-28&deg;C for two weeks. Continue to fertilize plants once a week with Miracle-Gro.</li>
 <li>For high-throughput screens, plant seeds directly in trays in sandy soil, cover the tray with plastic wrap until germination and maintain as above. After germination, add Osmocote slow release fertilizer and fertilize once a week with Miracle-Gro.</li>
</ol>
2. Cowpea Growth in Seedling Pouches
<ol>
 <li>Seeds are either germinated in a Petri dish, lined with several layers of Kimwipe paper, and transferred singly to pouches or placed directly into the paper groove of the pouch and germinated.</li>
 <li>Place the pouches in a plastic hanging file folder, two per folder, and arrange the folders in a rack in a vertical position. Place the rack in a controlled-environment chamber maintained at a temperature of 25-28 &deg;C and 16 h light / 8 h dark cycle. Racks also can be maintained in a greenhouse, but require a foil or stiff paper cover placed over the rack on both sides of the plant stems, to reduce potential for fungal contamination.</li>
 <li>Water the pouches once or twice per day with reverse osmosis water. About 10 to 14 days after seeding, when an adequate root system with tertiary root tips has developed (Figure 1), the seedlings are ready for inoculation.</li>
</ol>
3. Extraction of Nematode Eggs
Extraction of eggs from infected roots is modified from a protocol developed by Hussey and Barker (1973).
<ol>
 <li>Three to four days before nematode inoculation, extract root-knot nematode eggs from infected tomato roots. Before starting egg extraction, wash the work area thoroughly with hot water to avoid contamination. Also wash in hot water a blender, two buckets, a wire mesh support, a rubber mallet, three sieves of 425, 90 and 25 &mu;m aperture, a graduated cylinder and scissors.</li>
 <li>Stack the sieves from top to bottom in the following order: 425, 90 and 25 &mu;m aperture. The eggs will be collected on the 25 &mu;m sieve at the bottom. Put the sieves on a wire mesh supported in a sink. Put a bucket under the wire mesh to collect the run-through solution.</li>
 <li>Cut the tops of the infected plant(s), used as the source of the inoculum, and discard. Carefully remove the plant from the pot. Wash the roots by dipping in a plastic bucket full of water. Rinse the roots further under running water until the soil particles are washed away from the roots.</li>
 <li>Cut the roots with scissors into small pieces. Dispose of the taproot.</li>
 <li>Put the chopped roots from a single plant in a large plastic jar with a lid, add just enough 10% bleach solution to cover the roots and close the lid.</li>
 <li>Shake the jar containing the roots for 2 min.</li>
 <li>Open the jar and pour the roots onto the top sieve and wash with a hose attached to a misting nozzle. Wash well until all the bleach smell is gone. Use a mallet to tap the sides of the sieves to avoid clogging the sieve pores.</li>
 <li>Remove the top sieve and rinse the debris on the second sieve.</li>
 <li>Remove the second sieve and collect the fine debris, which includes the eggs, from the last sieve. With a gentle stream from a water bottle, move the debris to one side of the sieve. Collect the debris and eggs into a clean beaker with minimum amount of water.</li>
 <li>Sieve once more the water collected in the bucket through the 25 &mu;m sieve to collect any eggs that may have escaped. Rinse well with water and collect in the same beaker.</li>
 <li>Discard the plant debris from the top sieve and wash thoroughly all three sieves with pressurized water. Do not touch the mesh part of the sieves as it may distort the pore size.</li>
</ol>
4. Hatching Nematode Eggs
<ol>
 <li>Line a clean metal basket with a few layers of Kimwipe paper and fit on a glass Petri dish. Allow a 1-cm distance between the bottom of the basket and the dish.</li>
 <li>Pour the extracted nematode eggs onto the paper in the wire basket. Add enough liquid so that the bottom of the wire basket touches the water surface but is not immersed in water. Cover the top with a plastic lid.</li>
 <li>Every day check for water evaporation and add some water in the Petri dish so that the bottom of the basket touches the water. This will prevent the eggs from drying out.</li>
 <li>Every other day, collect the water that contains the J2s from the Petri dishes into a beaker. If not used immediately, aerate the collected inoculum at room temperature using a lab air supply or air generated by an aquarium pump. Use the aerated inoculum within 2 days. J2s may be collected from the hatching set up over a period of 6-8 days.</li>
</ol>
5. Tomato Root Infection in Pots or Trays and Evaluation of Infection
<ol>
 <li>Use three aliquots of the collected nematodes to count the number of J2s in a counting slide or dish, calculate the average and determine the desired volume for inoculation. Typically 3000 J2s are used per plant in pots and 500 J2s for plants grown in seedling trays.</li>
 <li>Stir the inoculum on a magnetic stirring plate at low speed.</li>
 <li>Before you inoculate the plants, make sure that the soil is moist but not too wet. For pot inoculation, make three holes of about half-pot depth in the sand around each tomato root system using a pencil (Figure 2). Inoculate each plant by delivering the J2s into the three holes using a pipette. Afterwards, cover the holes.</li>
 <li>For the high-throughput screens in trays, use a modified sealed tip needle with holes on the sides glued to a 5-ml pipette tip along a pipetter (Figure 3) to deliver the J2s into the soil. Alternatively, nematodes can be delivered as in step 5.3.</li>
 <li>Maintain the plants in a greenhouse at 24-27&deg;C for six to eight weeks. Continue to fertilize twice a month with Miracle-Gro.</li>
 <li>For evaluation of infection, carefully remove the plants from the pots and wash the roots (as described in step 3.3).</li>
 <li>Stain the egg masses blue by submerging the roots in 1 mg/L erioglaucine, for 15 min.</li>
 <li>Rinse the roots in water and evaluate by counting the stained egg masses on individual root system. An illuminated desk magnifier can be used to help visualize the egg masses.</li>
 <li>If the screened plants are needed for further genetic studies, do not cut the tops before washing the roots for evaluation. After staining and counting the egg masses, transplant the seedlings in organic soil, prune the tops heavily to reduce transpiration, and maintain in a greenhouse.</li>
</ol>
6. Cowpea Root Infection in Pouches and Evaluation of Infection
<ol>
 <li>Count the nematode inoculum and stir on a magnetic stirring plate as described earlier.</li>
 <li>Remove the pouches from the hanging folders and place on a horizontal surface. Inoculate each pouch with 1500 J2s in 5 ml. Lift the plastic cover of the pouch and distribute the nematodes evenly over the surface of the roots.</li>
 <li>Keep the pouches in a horizontal position for 24 h after inoculation, covered with dark paper to exclude light, then return to the hanging folders in the growth chamber.</li>
 <li>Water the plants as needed, once or twice daily, with half-strength Hoagland's solution (Hoagland &amp; Arnon, 1950; Table 1) until a response to the fertilizer is observed, typically enhanced foliage greening and more vigorous shoot growth. Usually the plants are watered with half-strength Hoagland 3 days in a row. Afterwards, keep the pouches moist with water.</li>
 <li>Approximately 30 days (range 28-35 days) after inoculation, infuse each pouch with about 10-20 ml of 75 mg/L erioglaucine. Keep the pouches flooded with the dye in a horizontal position overnight.</li>
 <li>Drain the pouches and evaluate the root systems by counting the egg masses under an illuminated desk magnifier.</li>
 <li>If the screened plants are needed for further genetic studies or breeding work, carefully pull out the roots from the paper, transplant in organic soil in pots and maintain in a greenhouse.</li>
</ol>
7. Representative Results
The appropriate stages of tomato and cowpea plants for nematode inoculations for the two systems described are shown in Figures 1 and 2. In addition, examples of well-infected roots of tomato and cowpea are shown in Figures 4 and 5. As in most disease resistance screens, it is advisable to use at least 6-10 plants per genotype for nematode inoculation to calculate the average of the infection rate. Variation in nematode infection between plants, of the same genotype, can be reduced by using uniform plant size and accurate amount and delivery of inoculum.
The use of trays and growth pouches enables screening of hundreds to thousands of plants in a small growth space. Growth pouches also allow fast and efficient non-destructive evaluation of root-knot nematode infections with no need for washing roots (Figure 4).
<img alt="Figure 1" src="/files/ftp_upload/3629/3629fig1.jpg" /> 
Figure 1. A two week-old cowpea plant grown in a pouch and ready for root-knot nematode inoculation.
<img alt="Figure 2" src="/files/ftp_upload/3629/3629fig2.jpg" /> 
Figure 2. A three week-old tomato plant ready for root-knot nematode inoculation.
<img alt="Figure 3" src="/files/ftp_upload/3629/3629fig3.jpg" /> 
Figure 3. A modified needle and pipette tip used for high-throughput inoculation of nematodes. The bottom of a needle was sealed and three sets of holes were drilled in the needle using a laser beam. Then, the modified needle was glued to a 5-ml pipette tip.
<img alt="Figure 4" src="/files/ftp_upload/3629/3629fig4.jpg" /> 
Figure 4. A tomato root system infected with root-knot nematodes.
<img alt="Figure 5" src="/files/ftp_upload/3629/3629fig5.jpg" /> 
Figure 5. A cowpea root system with egg masses stained with erioglaucine 30 days post-inoculation grown at 28 &deg;C.

<ol>
	<li>Bhattarai, K. K., Xie, Q. G., Mantelin, S., Bishnoi, U., Girke, T., Navarre, D. A., Kaloshian, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tomato+susceptibility+to+root-knot+nematodes+requires+an+intact+jasmonic+acid+signaling+pathway.">Tomato susceptibility to root-knot nematodes requires an intact jasmonic acid signaling pathway.</a> Mol. Plant Microbe Interact. 21, 1205-1214 (2008).</li><li>Ehlers, J. D., Matthews, W. C., Hall, A. E., Roberts, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inheritance+of+a+broad-based+form+of+root-knot+nematode+resistance+in+cowpea.">Inheritance of a broad-based form of root-knot nematode resistance in cowpea.</a> Crop Sci. 40, 611-618 (2000).</li><li>Hoagland, D. R., Arnon, D. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+water-culture+method+for+growing+plants+without+soil.">The water-culture method for growing plants without soil.</a> Calif. Agric. Exp. Stn. Circ. 347, (1950).</li><li>Hussey, R. S., Barker, K. 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+comparison+of+methods+of+collecting+inocula+of+Meloidogyne+spp.,+including+a+new+technique.">A comparison of methods of collecting inocula of Meloidogyne spp., including a new technique.</a> Plant Dis. Rep. 57, 1025-1028 (1973).</li><li>Martinez de Ilarduya, O., Moore, A. E., Kaloshian, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+tomato+Rme1+locus+is+required+for+Mi-1-mediated+resistance+to+root-knot+nematodes+and+the+potato+aphid.">The tomato Rme1 locus is required for Mi-1-mediated resistance to root-knot nematodes and the potato aphid.</a> Plant J. 27, 417-425 (2001).</li><li>Matkin, O. A., Chandler, P. A., Baker, K. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+U.C.-type+soil+mixes.+In:+The+U.C.+system+for+producing+healthy+container-grown+plants.">The U.C.-type soil mixes. In: The U.C. system for producing healthy container-grown plants.</a> Calif. Agric. Exp. Stn. Manual. 23, 68-85 (1957).</li><li>Omwega, C. O., Roberts, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inheritance+of+resistance+to+Meloidogyne+spp.+in+common+bean+and+the+genetic+basis+of+its+sensitivity+to+temperature.">Inheritance of resistance to Meloidogyne spp. in common bean and the genetic basis of its sensitivity to temperature.</a> Theor. Appl. Genet. 83, 720-726 (1992).</li><li>Omwega, C. O., Thomason, I. J., Roberts, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+nondestructive+technique+for+screening+bean+germplasm+for+resistance+to+Meloidogyne+incognita.">A nondestructive technique for screening bean germplasm for resistance to Meloidogyne incognita.</a> Plant Dis. 72, 970-972 (1988).</li><li>Perry, R. N., Moens, M., Starr, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Root-Knot Nematodes. , (2009).</li><li>Starr, J. L., Cook, R., Bridge, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Resistance to Parasitic Nematodes. , (2002).</li></ol>

We have nothing to disclose.

There are two critical steps for a successful nematode screen: preparation of a highly infective inoculum and using plants at the proper developmental stage. Hatching rate of root-knot nematode eggs is highly variable and ranges between 5- 50%. Therefore to obtain optimum levels of hatch and highly infective J2s, close attention must be paid to the egg extraction and hatching procedures. Eggs should be exposed to bleach for a minimum time period and bleach should be rinsed out well from the root mixture. When hatching the eggs, the slurry containing the eggs should not be submerged in water. In addition, avoid spilling the eggs into the underlying Petri dish where the hatched juveniles are collected. One way to avoid spilling the eggs during the hatching process is not to add water directly to the Kimwipe as the paper might break. Instead add the water directly to the Petri dish.
For best results, use freshly hatched J2s. If the hatch rate is low and more inoculum is needed, J2s can be stored at 15&deg;C for a longer period. Warm the stored inoculum to room temperature to revive the nematodes prior to inoculation. However, do not store the J2s for prolonged periods even at 15&deg;C as starved J2s will not infect efficiently.
Young seedlings are the best developmental stage for root-knot nematode inoculation. However, this needs to be balanced with the formation of an adequate root system providing sufficient numbers of root tips as points of entry for the J2s. Maintaining optimal plant growth condition is also critical for the screens. Avoid over-watering plants specially those grown in pouches. Over-watering the pouches, as indicated by standing water in the bottom of the pouch, can promote fungal growth and diminish root health.
With both assays further evaluation of nematode infection and calculation of the number of eggs/root system and eggs/gram of fresh root can be performed. For the tomato assays, individual roots are weighed and eggs extracted as described for inoculum preparation (section 3). When processing large number of plant samples, individual root systems could be macerated using a blender for egg extraction. The number of eggs should be counted in at least three aliquots and the eggs/gram of fresh root system calculated. For the pouch assays, the root system is pulled off the paper insert, weighed, and eggs extracted.

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.

Watch this Scientific Journal Video about High and Low Throughput Screens with Root-knot Nematodes Meloidogyne spp. at JoVE.com

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.

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 ...

high-low-throughput-screens-with-root-knot-nematodes-meloidogyne

Universidad de Cartagena UDEC

Research

JoVE Journal

Immunology and Infection

27.8K Views.  University of California, Riverside. 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.

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

Soil Sampling and Isolation of Entomopathogenic Nematodes (Steinernematidae, Heterorhabditidae)

Entomopathogenic nematodes (a.k.a. EPN) represent a group of soil-inhabiting nematodes that parasitize a wide range of insects. These nematodes belong to two families: Steinernematidae and Heterorhabditidae. Until now, more than 70 species have been described in the Steinernematidae and there are about 20 species in the Heterorhabditidae. The nematodes have a mutualistic partnership with Enterobacteriaceae bacteria and together they act as a potent insecticidal complex that kills a wide range of insect species.
Herein, we focus on the most common techniques considered for collecting EPN from soil. The second part of this presentation focuses on the insect-baiting technique, a widely used approach for the isolation of EPN from soil samples, and the modified White trap technique which is used for the recovery of these nematodes from infected insects. These methods and techniques are key steps for the successful establishment of EPN cultures in the laboratory and also form the basis for other bioassays that consider these nematodes as model organisms for research in other biological disciplines. The techniques shown in this presentation correspond to those performed and/or designed by members of S. P. Stock laboratory as well as those described by various authors.

Soil Sampling and Isolation of Entomopathogenic Nematodes Steinernematidae, Heterorhabditidae

Entomopathogenic nematodes are soil-inhabiting roundworms that parasitize a wide range of insects. We demonstrate sampling methods used for the isolation of these nematodes from the soil using two techniques: the insect baiting and the modified White trap, for the recovery of nematodes from soil samples and infected insect cadavers, respectively.

Entomopathogenic nematodes (a.k.a. EPN) represent a group of soil-inhabiting nematodes that parasitize a wide range of insects. These nematodes belong to ...

Environment

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.

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 ...

Genetics

Rapid Isolation of Wild Nematodes by Baermann Funnel

Beyond being robust experimental model organisms, Caenorhabditis elegans and its relatives are also real animals that live in nature. Studies of wild nematodes in their natural environments are valuable for understanding many aspects of biology, including the selective regimes in which distinctive genomic and phenotypic characters evolve, the genetic basis for complex trait variation, and the natural genetic diversity fundamental to all animal populations. This manuscript describes a simple and efficient method for extracting nematodes from their natural substrates, including rotting fruits, flowers, fungi, leaf litter, and soil. The Baermann funnel method, a classical nematology technique, selectively isolates active nematodes from their substrates. Because it recovers nearly all active worms from the sample, the Baermann funnel technique allows for the recovery of rare and slow-growing genotypes that co-occur with abundant and fast-growing genotypes, which might be missed in extraction methods that involve multiple generations of reproduction. The technique is also well suited to addressing metagenetic, population-genetic, and ecological questions. It captures the entire population in a sample simultaneously, allowing an unbiased view of the natural distribution of ages, sexes, and genotypes. The protocol allows for deployment at scale in the field, rapidly converting substrates into worm plates, and the authors have validated it through fieldwork on multiple continents.

This protocol outlines a method for efficiently extracting live nematodes from natural substrates in the field.

Beyond being robust experimental model organisms, Caenorhabditis elegans and its relatives are also real animals that live in nature. Studies of wild ...

Novel Metrics to Characterize Embryonic Elongation of the Nematode Caenorhabditis elegans

Dissecting the signaling pathways that control the alteration of morphogenic processes during embryonic development requires robust and sensitive metrics. Embryonic elongation of the nematode Caenorhabditis elegans is a late developmental stage consisting of the elongation of the embryo along its longitudinal axis. This developmental stage is controlled by intercellular communication between hypodermal cells and underlying body-wall muscles. These signaling mechanisms control the morphology of hypodermal cells by remodeling the cytoskeleton and the cell-cell junctions. Measurement of embryonic lethality and developmental arrest at larval stages as well as alteration of cytoskeleton and cell-cell adhesion structures in hypodermal and muscle cells are classical phenotypes that have been used for more than 25 years to dissect these signaling pathways. Recent studies required the development of novel metrics specifically targeting either early or late elongation and characterizing morphogenic defects along the antero-posterior axis of the embryo. Here, we provide detailed protocols enabling the accurate measurement of the length and the width of the elongating embryos as well as the length of synchronized larvae. These methods constitute useful tools to identify genes controlling elongation, to assess whether these genes control both early and late phases of this stage and are required evenly along the antero-posterior axis of the embryo.

Novel Metrics to Characterize Embryonic Elongation of the Nematode Caenorhabditis elegans

Here we detail protocols specifically designed to monitor morphogenic defects that occur during early and late phases of embryonic elongation of the nematode Caenorhabditis elegans. Ultimately, these protocols are designed to identify genes that regulate these phases and to characterize their differential requirements along the antero-posterior axis of the embryo.

Dissecting the signaling pathways that control the alteration of morphogenic processes during embryonic development requires robust and sensitive metrics. ...

Neuroscience

Visualizing Bacteria in Nematodes using Fluorescent Microscopy

Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on earth, nematodes and bacteria form a wide array of symbiotic associations that range from beneficial to pathogenic 1-3. One such association is the mutually beneficial relationship between Xenorhabdus bacteria and Steinernema nematodes, which has emerged as a model system of symbiosis 4. Steinernema nematodes are entomopathogenic, using their bacterial symbiont to kill insects 5. For transmission between insect hosts, the bacteria colonize the intestine of the nematode's infective juvenile stage 6-8. Recently, several other nematode species have been shown to utilize bacteria to kill insects 9-13, and investigations have begun examining the interactions between the nematodes and bacteria in these systems 9.
We describe a method for visualization of a bacterial symbiont within or on a nematode host, taking advantage of the optical transparency of nematodes when viewed by microscopy. The bacteria are engineered to express a fluorescent protein, allowing their visualization by fluorescence microscopy. Many plasmids are available that carry genes encoding proteins that fluoresce at different wavelengths (i.e. green or red), and conjugation of plasmids from a donor Escherichia coli strain into a recipient bacterial symbiont is successful for a broad range of bacteria. The methods described were developed to investigate the association between Steinernema carpocapsae and Xenorhabdus nematophila 14. Similar methods have been used to investigate other nematode-bacterium associations 9,15-18and the approach therefore is generally applicable.
The method allows characterization of bacterial presence and localization within nematodes at different stages of development, providing insights into the nature of the association and the process of colonization 14,16,19. Microscopic analysis reveals both colonization frequency within a population and localization of bacteria to host tissues 14,16,19-21. This is an advantage over other methods of monitoring bacteria within nematode populations, such as sonication 22or grinding 23, which can provide average levels of colonization, but may not, for example, discriminate populations with a high frequency of low symbiont loads from populations with a low frequency of high symbiont loads. Discriminating the frequency and load of colonizing bacteria can be especially important when screening or characterizing bacterial mutants for colonization phenotypes 21,24. Indeed, fluorescence microscopy has been used in high throughput screening of bacterial mutants for defects in colonization 17,18, and is less laborious than other methods, including sonication 22,25-27and individual nematode dissection 28,29.

To study the mutualism between Xenorhabdus bacteria and Steinernema nematodes, methods were developed to monitor bacterial presence and location within nematodes. The experimental approach, which can be applied to other systems, entails engineering bacteria to express the green fluorescent protein and visualizing, using fluorescence microscopy bacteria within the transparent nematode.

 
Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on ...

Biology

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 ...

High-Throughput Screening of Microbial Isolates with Impact on Caenorhabditis elegans Health

With its small size, short lifespan, and easy genetics, Caenorhabditis elegans offers a convenient platform to study the impact of microbial isolates on host physiology. It also fluoresces in blue when dying, providing a convenient means of pinpointing death. This property has been exploited to develop high-throughput label-free C. elegans survival assays (LFASS). These involve time-lapse fluorescence recording of worm populations set in multiwell plates, from which population median time of death can be derived. The present study adopts the LFASS approach to screen multiple microbial isolates at once for the effects on C. elegans susceptibility to severe heat and oxidative stresses. Such microbial screening pipeline, which can notably be used to prescreen probiotics, using severe stress resistance as a proxy for host health is reported here. The protocol describes how to grow both C. elegans gut microbiota isolate collections and synchronous worm populations in multiwell arrays before combining them for the assays. The example provided covers the testing of 47 bacterial isolates and one control strain on two worm strains, in two stress assays in parallel. However, the approach pipeline is readily scalable and applicable to the screening of many other modalities. Thus, it provides a versatile setup to rapidly survey a multiparametric landscape of biological and biochemical conditions that impact C. elegans health.

High-Throughput Screening of Microbial Isolates with Impact on Caenorhabditis elegans Health

Gut microbes may positively or negatively impact the health of their host via specific or conserved mechanisms. Caenorhabditis elegans is a convenient platform to screen for such microbes. The present protocol describes high-throughput screening of 48 bacterial isolates for impact on nematode stress resistance, used as a proxy for worm health.

With its small size, short lifespan, and easy genetics, Caenorhabditis elegans offers a convenient platform to study the impact of microbial isolates on ...

In vivo and In vitro Infection of Potato Roots with Plant Parasitic Nematodes for the Assessment of Induced Structural Changes

Soil-dwelling plant parasitic nematodes (PPNs) are important potato pests that cause lesions and/or change plant roots structure, leading to reduced crop fitness and productivity. Research on the cellular and subcellular mechanisms of PPNs infection and development can resort to field plants or seedlings under greenhouse conditions. Field studies are more representative of natural environments but are subjected to the unpredictability of environmental conditions that can heavily influence research outcomes. Greenhouse studies allow higher control over environmental variables and higher safety against contaminants or pathogens. However, in some hosts, genetic diversity becomes an important factor of variability and influences the host-parasite complex response. We have developed in vitro co-cultures of transgenic roots with PPNs as a reliable alternative that occupies less space, requires less time to obtain, and is free from contamination or from host genetic variability. Co-cultures are obtained by introducing aseptic PPNs to host in vitro transgenic roots. They can be maintained indefinitely, which makes them excellent support for keeping collections of reference PPNs. In the present work, a protocol is detailed for the controlled infection of in vivo potato roots with the root lesion nematode and for establishing in vitro co-cultures of potato transgenic roots with the root-knot nematode. The in vitro co-cultures provided a laboratory proxy for the natural potato infection condition and produced nematode life stages irrespective of season or climate conditions. Additionally, the methodology used for structural analysis is detailed using histochemistry and optical microscopy. The acid fuchsin dye is used to follow nematode attack sites on roots, while differential staining with Periodic acid-Schiff (PAS) and toluidine blue O highlights nematode structures in potato internal root tissue.

The protocol describes infection of Solanum tuberosum roots with plant parasitic nematodes under in vivo greenhouse conditions and potato in vitro transgenic roots for histochemical analysis of root structure through optical microscopy.

Soil-dwelling plant parasitic nematodes (PPNs) are important potato pests that cause lesions and/or change plant roots structure, leading to reduced crop ...

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 ...

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Nematode Synchronization: A Method to Obtain Populations of Worms in Identical Stages of Development

Source: Sarasija, S. and Norman, K. R. <a href="https://jove.unicartagenaproxy.elogim.com/video/59277/measurement-oxygen-consumption-rates-intact-caenorhabditis">Measurement of Oxygen Consumption Rates in Intact&#160;Caenorhabditis elegans</a>.&#160;J. Vis. Exp. (2019).
In this video, we show a method of producing a developmentally synchronized population of sterile adult C. elegans. The example protocol synchronizes worms to study oxygen consumption rates in live animals.

Synchronizacja nicieni: metoda uzyskiwania populacji robaków w identycznych stadiach rozwoju

Source: Sarasija, S. and Norman, K. R. Measurement of Oxygen Consumption Rates in Intact&#160;Caenorhabditis elegans.&#160;J. Vis. Exp. (2019).
In this ...

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

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Rozdziały w tym wideo

Soil Sampling and Isolation of Entomopathogenic Nematodes (Steinernematidae, Heterorhabditidae)

Screening Cotton Genotypes for Reniform Nematode Resistance

Rapid Isolation of Wild Nematodes by Baermann Funnel

Novel Metrics to Characterize Embryonic Elongation of the Nematode Caenorhabditis elegans

Visualizing Bacteria in Nematodes using Fluorescent Microscopy

Culturing and Screening the Plant Parasitic Nematode Ditylenchus dipsaci

High-Throughput Screening of Microbial Isolates with Impact on Caenorhabditis elegans Health

In vivo and In vitro Infection of Potato Roots with Plant Parasitic Nematodes for the Assessment of Induced Structural Changes

Genetic Screens

Synchronizacja nicieni: metoda uzyskiwania populacji robaków w identycznych stadiach rozwoju