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

Summary

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.

Abstract

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.

Introduction

Root and tuber crops rank 4^th among the world's most important staple foods. Potato (Solanum tuberosum L.) is one of the most important cultivated tubers. It had its origin in the Andes mountains of South America, but after being introduced to Europe in the 16^th century quickly became the most common food source for the population with a lower income. Today, potatoes make up 1.7% of the world's caloric intake¹. Crop production is heavily affected by plant pests and pathogens, of which plant parasitic nematodes (PPNs) can cause average yield losses that rise up to 12%². Plant parasitic nematodes are responsible for some of the most damaging diseases to crops in modern agriculture. Soil-dwelling PPNs impose heavy losses to farmers because they affect plant roots and interfere with crop productivity by reducing production and/or injuring products, turning them unmarketable³. These dangerous phytoparasites use their stylet (a needle-like mouthpart) to puncture root cells and feed on cell content. Some PPNs feed from outside the roots, others enter the root and cause tissue damage (migratory), while others enter the roots and become sedentary, heavily changing root structure to facilitate feeding⁴. The main PPNs affecting potato are the potato cyst nematodes, Globodera spp., root-knot nematodes (RKN), Meloidogyne spp., root lesion nematodes, Pratylenchus spp., the false root-knot nematode Nacobbus aberrans, and the potato rot nematode Ditylenchus destructor. For these PPNs, different feeding habits induce different structural changes in host root tissues^5,6. Research on the mechanisms of PPN infection and host response is often performed through field or greenhouse trials to maintain reference PPN culture collections or to perform large scale experiments^7,8. Testing under natural conditions is strongly influenced by environmental variation and biotic or abiotic stress factors. Greenhouse bioassays are a closer alternative to a natural condition while allowing a relative control of environmental variation and limiting the influence of abiotic and biotic stress. However, host genetic diversity can still be a challenge for trials that require a finer control of biological variability. These limitations can be overcome by resorting to in vitro plant tissue cultures. These are versatile laboratory systems with many advantages for PPNs disease research. For soil-dwelling PPNs, in vitro cultures of transgenic roots are a useful tool for research in laboratory conditions^9,10.

Transgenic roots, or hairy roots (HR), are obtained after infection of plant material with Rhizobium rhizogenes (Riker et al. 1930) Young et al. 2001¹¹. This gram-negative bacterium induces the transfection of its Ri plasmid into the host genome and changes the regulation of plant hormone biosynthesis, promoting the formation of root tissue¹². Transgenic roots can be maintained indefinitely under asepsis in a culture medium. The advantages of using HR for studying PPNs are a high growth rate in the absence of plant growth regulators that influence nematode infection and development, a high ratio of biomass production per unit time, and cellular integrity and longevity, which determine a higher genetic and biochemical stability⁶. By resorting to in vitro transgenic roots, PPNs genotypes can be maintained indefinitely under laboratory conditions, infection and PPNs development can be easily followed, host genetic variability can be reduced, manipulation of host molecular makeup can be directly linked to nematode response, and host and parasite structural changes can be more accurately followed^6,13. For studies on PPN diseases of potato, in vitro transgenic root co-cultures allow carrying out experiments independently of season or potato tuber dormancy.

In this protocol, the traditional methodology of PPNs maintenance and in vivo infection of potato plants are detailed. For the structural analysis of infected roots, an improved methodology based on the establishment of in vitro co-cultures of transgenic potato roots with PPNs is also detailed as an alternative that allows a higher control of environmental and host genetic variability. To follow PPNs infection and development in the root tissue, histochemistry is employed to aid in PPNs observation under optical microscopy. The overall aim of this protocol is to optimize the study of PPN-host interactions, ensuring more controlled and reproducible conditions for experimentation while facilitating detailed structural and developmental analyses of nematodes in the root tissue.

Protocol

1. Infection of greenhouse-grown potato plants

NOTE: Greenhouse trials are performed with suspensions of PPNs in mixed life stages or second-stage juveniles (J2), depending on the specific life cycle of the PPN pest. For this protocol, suspensions of mixed life stages of the root lesion nematode (RLN) Pratylenchus penetrans were used. PPNs can either be reared in the lab or requested from certified reference laboratories.

1. Multiplication and maintenance of root lesion nematodes
   NOTE: Sterilized carrot disks are used for the multiplication and maintenance of RLNs¹⁴. Use commercially acquired carrots (var. Nice) with no visible damage to reduce microbial contamination. Preferably, they should be free from pesticides to avoid hindering RLN development.
   1. Wash the carrot in running tap water to remove larger debris, and afterward, with a common detergent solution (1 drop per 40 mL of water) to remove finer debris. Dry with laboratory paper towels.
   2. Under asepsis, in a vertical flow hood, insert a sterilized metal skewer on the top of the carrot (1 to 2 cm inwards) so it can be more easily held.
   3. With the help of a wash bottle with a nozzle, wet the carrot with 96% (v/v) ethanol. Blot the bottom tip of the carrot in a sterilized filter paper and carefully take it to the flame.
      CAUTION: Be aware that ethanol strongly ignites, so stand at a distance.
   4. Peel the carrot from the top down using a sterile peeler and then repeat the previous step. Discard the top and bottom sections (2 cm inwards) and set the middle section of the carrot in a sterile Petri dish (150 mm in diameter). Using a sterile blade and tweezers, carefully cut 1 cm thick sections from the approximately 2 cm diameter portion of the carrot (Figure 1).
   5. Transfer the sections to sterile Petri dishes (60 mm diameter) and seal the border with transparent film. Using a UV light, sterilize the surface of the carrot disks for 60 min on each side.
   6. Keep at 25 °C for 1-2 weeks in darkness and discard any carrot disks that begin to show signs of microbial contamination¹⁵.
      NOTE: Visible signs of contamination are excessive browning (rotting), liquid accumulation in the lower border of the carrot disk, or fungal mycelium growth on the surface.
   7. The remaining carrot disks are ready to be infected with RLN suspensions. Begin by making an X-shaped incision at the center of the carrot disk using a sterile blade. Be sure to cut only halfway deep.
   8. Inoculate the RLN by pipetting 50 µL of a suspension containing at least 50 mixed life stages in the center of the X-shaped wound. Close the Petri dish and seal the border with transparent film to avoid desiccation.
      1. Determine the average number of nematodes in the suspension by counting five 50 µL aliquots under a binocular stereomicroscope (40x) at room temperature in a concave slide. Set the suspension of mixed life-stage RLNs to 1000 per mL by adding water to the suspension or waiting for nematodes to settle (approximately 60 min) and lowering the volume by decanting the surface water.
   9. Keep the carrot disks at 25 °C, in darkness, for up to 3 months and follow weekly under a binocular stereomicroscope for signs of necrotic lesions, resultant of RLN population growth.
      NOTE: Successfully parasitized carrot discs can be stored at 11 °C for later use for up to 2 months but check regularly for microbial contamination. Parasitized carrot discs that show signs of microbial infection must be decontaminated by autoclaving before being discarded.
   10. Under the flow hood, extract RLNs by transferring carrot disks with visible tissue necrosis at the inoculation site (Figure 1) to an 8 cm diameter and 75 µm mesh sieve set in a sterile glass bowl. Leave a small 1 cm gap between the bottom of the sieve and the bowl concavity to collect the RLNs.
       NOTE: In the lack of a commercially acquired sieve, one can be crafted from an 8 cm diameter plastic tube/a sturdy plastic cup, and a tight mesh gauze. Use rubber bands to secure the gauze to the plastic tube or cup.
   11. Pour an antibiotic solution into the sieve until the carrot disks are covered and keep for 12 h (overnight) in darkness. The RLNs egress from the carrot disks and settle at the bottom of the bowl. The antibiotics solution should be prepared extemporaneously with the extraction by adding 50 µg/mL each of kanamycin and carbenicillin in sterilized distilled water¹⁶.
       NOTE: Antibiotics stock solutions are prepared at 50 mg/mL by dissolving 0.5 g kanamycin monosulfate or 0.5 g carbenicillin disodium each in 10 mL of sterilized distilled water. Stock solutions are filtered (0.22 µm mesh) in the flow hood and can be kept at -20 °C for up to 1 year.
   12. Remove the sieve, use a sterilized glass pipette to draw the RLNs from the bottom of the bowl into a sterilized glass staining block (4 cm x 4 cm x 1 cm), and wash by pipetting 1 mL of antibiotic solution. Wait 30 to 40 min for the nematodes to settle before collecting the used antibiotic solution. Repeat this wash 4x-5x.
   13. Use the aqueous suspension with the RLNs immediately or keep it at 11 °C for a longer storage period (up to 2 months).
2. In vivo infection of potato plants with PPNs
   NOTE: To grow susceptible potato plants (S. tuberosum var. Désirée), certified seed potatoes should be acquired from agro-dealers between January and March. Choose certified seed potatoes because they are sold with a phytosanitary passport, ensuring they are uncontaminated with quarantine phytoparasites. As a precaution, an initial step of disinfection with a 10% bleach solution followed by washing in running tap water can be performed to ensure disinfection of the potato tuber surface. Common commercialized potatoes are not recommended since the treatments imposed to reduce sprouting and vigor may interfere with potato growth and response to infection.
   1. Select same-sized potato tubers and discard those with holes, bruises, or softer sections. Gently remove all grown sprouts (1 mm) before sowing to synchronize sprouting.
      NOTE: If needed, store the seed potatoes in a well-ventilated, dry, and dark place before sowing.
   2. Fill 5 L pots (22 cm x 18 cm) with a 1:1 mixture of autoclaved soil and fine coarse sand mixed with 22.5 g of slow-release NPK fertilizer (12-12-12) and sow the potatoes at 9 cm below the soil surface.
      NOTE: Soil and sand should be sieved to remove debris larger than 2 mm, autoclaved 2x at 121 °C for 15 min, and dried at 100 °C for 1 to 2 days, with frequent mixing. Aerate the next 7 – 10 days by mixing frequently, before use.
   3. Keep the pots in a greenhouse under humid conditions (50%-70% humidity) and water frequently (keep the soil at 70% maximum water holding capacity), avoiding temperature extremes, until potato plant shoots begin emerging at the soil surface.
   4. After plant emergence, use freshly extracted RLN suspensions to infect the potato roots. Begin by creating evenly distributed 4 to 6 holes (1 cm wide) around the plant to seed depth.
   5. Evenly pipette an 8 mL suspension of 30,000 living mixed life stage RLNs into the holes, so that the inoculum is at a ratio of 4 live RLNs per g of soil mixture, and cover with soil mixture. For the pots with RLNs, withhold watering on the day of inoculation.
      NOTE: RLNs are counted under a stereomicroscope (40x). Dead nematodes are nonmotile and have an extended shape, while live nematodes are generally moving (non-extended shape). Physical prodding is used to ascertain mortality.
   6. Keep the pots for 2 months under the conditions described above (Figure 2). Afterward, uproot the potato plants and weigh the shoots and roots separately.
   7. Carefully wash the root system before checking the location of RLN attack sites through staining techniques⁵.

2. Establishment of in vitro co-cultures of potato transgenic roots with PPNs

1. Establishing in vitro potato transgenic roots
   NOTE: For this protocol, we used Rhizobium rhizogenes carrying the gus reporter gene co-integrated in the Ri plasmid and driven by a double 35S promoter (A4pRiA4::70GUS)¹⁷. Bacteria can be acquired from commercial sources or requested from certified reference laboratories.
   1. To obtain bacteria at the exponential growth phase, spread plate R. rhizogenes in a Luria-Bertani (LB)¹⁸ solid medium plate and keep overnight at 26 °C.
      NOTE: LB medium can be acquired commercially or prepared in the laboratory by adding 10 g/L peptone, 5 g/L yeast extract, 10 g/L NaCl, and 15 g/L agar and then steam sterilizing for 15 min at 121 °C.
   2. With an inoculation loop, pick a colony and inoculate 10 mL of liquid LB broth (LB medium without agar) in a sterile 50 mL flask. Keep overnight in the dark at 26 °C under agitation (180 rpm).
   3. Measure the absorbance of the liquid culture until A₆₀₀ reaches 0.6. At this stage, bacteria are at the exponential growth phase and are used for inoculating the plant material.
   4. Inoculation is performed in aseptic fresh potato tubers. To sterilize the surface of potato tubers, begin by washing in running tap water to remove larger debris, and then with a common detergent solution (1 drop per 40 mL of water), with vigorous agitation, to remove the finer debris.
      NOTE: Select a potato variety with known susceptibility to the PPNs in use. For this protocol, we used S. tuberosum var. Désirée.
   5. Place the tubers in a container, cover with a commercial bleach solution (1:4, commercial bleach in tap water) and close. Mix for 15 min, dispose of the bleach solution and rinse 3x with sterilized tap water.
   6. In a flow hood, immerse the tubers in an ethanol solution (80%, v/v) for 15 min with vigorous agitation, dispose of the ethanol and rinse 3x with sterilized tap water.
   7. Using a sterile scalpel, remove the peripheral portions of the tubers (approximately 50% of the tuber from the surface inwards), and section the inner central piece into 0.5 cm thick segments¹⁹. Inoculate immediately with the bacterial suspension prepared in step 2.1.3.
   8. For inoculation, dilute the bacterial suspension by adding 1 mL of bacterial suspension to 9 mL of Schenk and Hildebrandt²⁰ (SH) medium supplemented with 30 g/L sucrose at pH = 5.6. Dip the tip of a sterile scalpel in the diluted suspension and wound the surface of the potato segment. Repeat this step 5x for each potato segment.
   9. Dry the segments of excess humidity in sterile filter paper for 1 min, place them in semi-solid SH medium (SH medium with 30 g/L sucrose, 8 g/L agar, pH = 5.6) and keep in darkness at 25 °C for plasmid transfection to occur.
   10. After 3 days, transfer the infected segments to plates of semi-solid SH medium supplemented with 150 µg/mL of each of the antibiotic's cefotaxime and carbenicillin. Keep for over 3 months with weekly medium renewal to ensure elimination of the bacteria.
       NOTE: Cefotaxime stock solution can be prepared at 100 mg/mL by dissolving 1 g cefotaxime sodium in 10 mL of sterile demineralized water and filtering (0.22 µm mesh) under the flow hood. Antibiotics can be kept at -20 °C for up to 1 year.
   11. After 3 months, transgenic root growth is extensive. Transfer the roots to a fresh, semi-solid SH medium without antibiotics by gathering a 1 g root clump with the tips of a sterile tweezers and placing it at the center of the culture medium in a new plate (Figure 3).
       NOTE: Approximately 1 month after infection, small masses of cell growth appear in the surface of the potato segment, from where the transgenic roots start developing⁶. Be sure to keep them in contact with the culture medium otherwise they might desiccate.
   12. To ensure genetic and metabolic stability, keep transgenic roots under a monthly sub-culture routine (as described in step 2.1.11.) at 25 °C in darkness for over 1 year before infecting with PPNs.
       NOTE: Ensure that at least six replicates are kept at every step of the protocol since unwanted microbial contamination often occurs. Once established, a single co-culture plate can be used as an inoculum for several new co-cultures; however, be sure to keep at least six replicate plates.
2. Establishing in vitro co-cultures of transgenic potato roots with PPNs
   NOTE: To obtain co-cultures of transgenic roots with PPNs, the nematode sterilization process is critical. For the present protocol we used second stage juveniles of the quarantine root-knot nematode Meloidogyne chitwoodi. Nematode inoculum can be obtained from certified reference laboratories in the form of root galls.
   1. Under a binocular stereomicroscope (20x), isolate nematode egg masses from the root galls with sterile ultra-fine point tweezers. Place egg masses in a covered Petri dish with 5 mL of sterile tap water and let the eggs hatch for 48 h. Set the J2 suspension to 100 nematodes per mL.
   2. In a flow hood, pipette 5 mL of a suspension containing 500 J2 into a sterile 20 µm mesh sterile sieve and wash with sterile tap water.
   3. Immerse the bottom half of the sieve containing the J2s in a 20 % hydrogen peroxide (H₂O₂) solution, and mix manually in a circular motion for 15 min.
   4. Wash the sterile J2s by dispensing sterile tap water through the sieve. Repeat this step 3x. In the final wash, tilt the sieve so that the nematodes collect at the sieve border. Recover the sterile nematode suspension by pipetting 1 mL of sterile ultrapure water in the sieve border, and store at 11 °C or use immediately.
      NOTE: Success of sterilization can be assessed by plating a 100 µL aliquot of the nematode suspension in SH medium and regularly monitoring, for 1 week, for contamination.
   5. In the flow hood, subculture a 1 g clump of potato transgenic roots (as described in step 2.1.11.) onto SH plates with 100 sterile nematodes (100 µL of a suspension with 1000 J2s per mL). After 2 to 3 weeks, small galls begin to appear in the new roots.
   6. Follow the co-culture regularly under an inverted microscope (100x), and when egg masses begin to be noticeable, subculture to a new semi-solid SH medium plate, making sure galls are taken with the root clump (Figure 4). Keep co-cultures under a monthly sub-culture routine at 25 °C in darkness.

3. Structural analysis of PPNs infection

NOTE: To follow PPNs induced changes in root tissue structure, histochemical staining techniques are used to contrast tissues with different chemical compositions. Differential staining is performed in roots masses or in thin sections of fixed root material, where specific dyes react with the target tissue according to their chemical affinity²¹. For the present protocol, we used acid fuchsin, or periodic acid-Schiff's reagent (PAS) combined with toluidine blue O dyes for differential staining.

1. Plant parasitic nematodes distribution in roots stained with acid fuchsin
   NOTE: To follow the distribution of PPNs throughout the root system, acid fuchsin is used to stain the nematode muscle tissue in a red hue⁵.
   1. Begin by washing the root system under running tap water for 5 min to remove any soil debris (in vivo plant roots) or residues of the culture medium (in vitro transgenic roots). Use your fingers in a circular motion to aid in detaching the soil from the root system.
   2. Cut the root system into 1 to 2 cm long sections and place it inside a 150 mL beaker. Dispense 70 mL of a 1.5 % sodium hypochlorite (NaOCl) solution and mix vigorously for 4 min to clear root tissues. Afterward, dispose of the NaOCl solution, rinse the roots in running tap water, and soak for 15 min in demineralized water to remove residual NaOCl²².
      NOTE: Chlorine bleach contains a minimum of 5.25% NaOCl, so add 20 mL of chlorine bleach to 50 mL of water to obtain a 1.5% NaOCl solution. For softer material (e.g., softer tissue of young roots or of in vitro grown transgenic roots) use a 0.9% NaOCl solution, while for harder material (older lignified roots) a 2.0% NaOCl solution is used.
   3. Drain the cleared roots and set them on a borosilicate glass beaker with 30 mL of demineralized water. Pipette 1 mL of acid fuchsin stain solution, mix manually and boil for 30 s on a hot plate. Let the glass beaker cool down, drain the solution and wash the stained roots in running tap water.
      NOTE: The acid fuchsin stain solution is made by dissolving 3.5 g of acid fuchsin dye powder in 250 mL of acetic acid and adding 750 mL of demineralized water. Staining is always followed by a destaining step to remove unused excess stains and enhance specimen contrast.
   4. Destain by adding 10-30 mL of glycerin acidified with a few drops of HCl (5N)¹³.
   5. Observe under the stereomicroscope or inverted microscope to roughly assess where in the root structure the PPNs are preferentially attacking (Figure 5 and Figure 6).
2. Assessment of root cell morphology with periodic acid-Schiff (PAS)/toluidine blue O
   NOTE: The influence of PPNs infection or damage to root cell morphology can be assessed under the microscope by first fixating, sectioning, and differentially staining infected potato roots.
   1. In a closed sample vial, fix fresh root material with 2.5% glutaraldehyde, prepared in 0.1 M sodium phosphate buffer, at pH 7.2, for 24-48 h, at room temperature⁶.
      CAUTION: Glutaraldehyde is toxic. Avoid inhalation and contact. Use protective lab coat and gloves and work in a fume hood. Sample vials should be closed unless in rinsing or in vacuum procedures. Dispose of glutaraldehyde following Hazardous Waste Procedures rules. Steps 3.2.1 to 3.2.5 are performed in a fume hood to avoid reagents inhalation.
      NOTE: To prepare 1 L of sodium phosphate buffer, pH 7.2, add 68.4 mL of 1M Na₂HPO₄ (141.96 g in 1 L solution) to 31.6 mL of 1M NaH₂PO₄ (119.98 g in 1 L solution) and fill up to 1 L by adding 900 mL of demineralized water. To help infiltration of the fixative, place the uncapped sample vials with root material under a low vacuum (26 mm Hg) for 2 min in a desiccator connected to a vacuum pump.
   2. With a glass Pasteur pipette, discard the fixative solution and wash fixated roots with sodium phosphate buffer (3x).
   3. Begin gradually dehydrating the fixated root tissue by replacing the buffer with a 10 % ethanol solution (v/v) for 15 min. Afterward, exchange with a 20% ethanol solution using a glass pipette, and keep the roots imbedded for 15 min. Continue with the graded succession of increasing ethanol concentrations (30%, 40%, 50%, 60%, 70%, 80%, and 90% for 15 min each) until the pure ethanol step, where the roots should be kept for 1 h.
   4. Embed the dehydrated roots gradually in resin (2-hydroxyethyl methacrylate). With a glass pipette, replace the pure ethanol with a 3:1 (v/v) ethanol/resin solution and keep for 24 h at 4 °C. Follow this with 1:1 and 1:3 (v/v) ethanol/resin solutions, each with a 24 h incubation period at 4 °C. Afterwards replace the 1:3 (v/v) solution with pure resin supplemented with dibenzoyl peroxide (1%) as a polymerization initiator.
   5. Set the samples in a resin mold tray, add a resin:dimethyl sulfoxide 15:1 (v/v) mixture, and keep at 60 °C over a hot plate for 48 h for the resin to harden.
   6. Set the impregnated samples in a rotary microtome equipped with a tungsten knife and slice 2-5 µm sections onto glass slides, according to the manufacturer's instructions.
   7. Begin the differential staining by immersing the slides for 10 min in a glass staining jar with a 15% 2,4-dinitrophenylhydrazine solution in acetic acid, at room temperature. Afterwards wash carefully under running tap water for 15 min and dry in an oven at 60 °C (15 min).
   8. Following, immerse the slides in periodic acid (1%) for 10 min, and then wash under running tap water for 5 min and leave to dry in an oven at 60 °C (15 min).
   9. Immerse the slides in Schiff′s Reagent (composed of 1% pararosaniline and 4% sodium metabisulfite, in 0.25 M hydrochloric acid), for 30 min. Afterwards, wash with a sodium metabisulfite solution (0.5%) in hydrochloric acid (0.05 M), for 2 min. Repeat 3x. Finally wash in running tap water for 5 min, and dry at room temperature.
   10. For contrast, stain by immersing the slides in 0.05% toluidine blue O for 15 min, washing in running tap water for 15 min and drying in an oven at 60 °C (15 min).
   11. Observe under a microscope (100x) equipped with image capture hardware (Figure 7).

Results

Carrot disks can be used to multiply and maintain several types of migratory PPNs²³. For the RLN, this technique is generally used to maintain reference collections of nematode species or isolates. Using carrot disks, an average 100x increase in nematode populations can be obtained in a period of 3 months (Figure 1). However, nematode numbers vary widely (between 30x and 200x), mainly owing to nematode genetic diversity and/or variation in nutritional contents of carr...

Discussion

The study of the mechanisms of infection and disease development in plants attacked by soil-dwelling PPNs is difficult because these phytoparasites generally infect the inner tissues of the root system and induce unspecific symptoms in the shoots. Despite the controlled environmental conditions of the greenhouse, sprouting potato tubers and the growth of potato plants are still favored in the spring and summer months, reducing the experimental period available to one season per year. Also, a substantial number of pots ha...

Materials

Name	Company	Catalog Number	Comments
2,4-Dinitrophenylhydrazine	Sigma-Aldrich	D199303
2-Hydroxyethyl methacrylate	Sigma-Aldrich	17348
Acetic acid	Sigma-Aldrich	695092
Acid Fuchsin	Sigma-Aldrich	F8129
Benzoyl peroxide	Sigma-Aldrich	B5907
borosilicate glass beaker	Sigma-Aldrich	Z231827
Carbenicillin disodium salt	Sigma-Aldrich	C3416
Cefotaxime sodium salt	Sigma-Aldrich	C7039
Dimethyl sulfoxide	Sigma-Aldrich	472301
Ethanol	Supelco	1.00983
Fertilizer	Compo Expert
Flower pot 5 L	VWR	470049-676
Glutaraldehyde	Sigma-Aldrich	354400
Glycerol	Sigma-Aldrich	G7893
Hydrochloric acid	Sigma-Aldrich	258148
Kanamycin monosulfate	Sigma-Aldrich	BP861
LB Broth with agar	Sigma-Aldrich	L3147
MCE syringe filter	Millipore	SLGSR33SS
PARAFILM M sealing film	BRAND	HS234526B-1EA
Pararosaniline hydrochloride	Sigma-Aldrich	P3750
Periodic acid	Sigma-Aldrich	P0430
Phyto agar	Duchefa Biochemie	P1003
Scalpel blade no. 24	Romed Holland	BLADE24
Schenk & Hildebrandt Basal salt medium	Duchefa Biochemie	S0225
Schenk & Hildebrandt vitamin mixture	Duchefa Biochemie	S0411
Schiff′s reagent	Sigma-Aldrich	1.09033
Sodium metabisulfite	Sigma-Aldrich	161519
Sodium phosphate dibasic	Sigma-Aldrich	S9763
Sodium phosphate monobasic	Sigma-Aldrich	S5011
Soil / Substrate	Compo Sana
Stainless Steel Tweezers	Sigma-Aldrich	22435-U
Sucrose	Duchefa Biochemie	S0809
Toluidine Blue O	Sigma-Aldrich	198161