Steinernematids and Heterohabditids

Culture, Storage and Transport

Relatively efficient and economical methods for large-scale culturing of steinernematids and heterorhabditids are available. Workable procedures for storage and transport of the nematodes have been developed. Moreover, recent evidence indicates that these nematodes can be stored anhydrobiotically, which will increase the feasibility of using the nematodes in integrated pest management systems.

General Considerations in Nematode Culture

When determining if and how to culture steinernematids and heterorhabditids, many factors must be taken into consideration.

  1. What facilities and manpower are available for the task? The requirements for each option will become apparent as they are described.
  2. How many nematodes are required? In vivo culture is most appropriate for maintenance of colonies and production of lJs for laboratory and small-scale field tests. For large-scale production in vitro methods are the most practical.
  3. Which nematodes are needed, and how long will they be stored before use? Steinernematids have much better storage qualities than heterorhabditids. In vivo production of large numbers of heterorhabditids within a short enough time interval to avoid long storage periods may be difficult.

In Vivo Production

Since steinernematids and heterorhabditids infect and reproduce in a broad spectrum of insects, they may be readily reared in vivo in the laboratory. Galleria mellonella is often used as a host because it is widely available, easily reared, very susceptible and an excellent host for nematode reproduction. Up to 200,000 S. feltiae (Dutky et al. 1964) and 350,000 H. bacteriophora infectives (Milstead and Poinar 1978) have been harvested from one last-instar Galleria. Average production is much less, on the order of 30,000 to 50,000 lJs per insect. Rearing procedures and sources for Galleria are found in Appendix 1, No.,2.

The basic in vivo production method is outlined below; Poinar (1979) describes and/or references various modifications.

1. Infecting Galleria: Let IJ suspension warm to room temperature (20-24 C). Examine the nematodes briefly under dissecting microscope. Dead dauers will generally be straight. Live dauers will actively move about. Dilute 1-ml suspension in an appropriate quantity of sterile distilled water (sdw) to yield a suspension near 200 nematodes/ml. Count the lJs (see "Quantification of Nematodes and Bacteria" in the "Laboratory Experimentation" section, below). Adjust the suspension to 200 nematodes/ml. If there are fewer than 200 nematodes/ml, either start over or let the lJs settle and remove the appropriate amount of supernatant.

Evenly distribute 1 ml of the nematode suspension on a 9.0-cm Whatman #1 filter paper in the lid of a 100 x 14 mm plastic petri dish. Add 10 conditioned Galleria larvae (Appendix 1, No. 2). The goal is to have about 20 nematodes per larva. Too many nematodes per larva produce few progeny due to competition and/or contamination with foreign bacteria. Cover the lid (containing nematodes and Galleria) with the inverted petri dish bottom. Label the petri dishes and store them in a plastic bag (to conserve moisture) at room temperature. Place infected larvae into White traps (Vvhite 1927) 5-7 days after infection. Steinernema-infected larvae will be yellowish brown and limp when held with forceps. Heterorhabditis-infected larvae turn brick-red and are also limp. Heavily contaminated insects or insects dying of other causes are usually blackish and smell putrid. It is best not to harvest from them as their production is poor and they contaminate the whole batch.

2. Harvesting: To make White traps (Vvhite 1927, Fig.2), place a 9.0-cm Whatman #1 filter paper in a concave-side-up watch glass in a large glass petri dish (150 x 20 mm). Autoclave for 20 minutes at 121 C. Pour about 70 ml sdw or 0.1% formalin into the petri dish. Do not put any water into the watch glass. Drape the filter paper over the watch glass so that it comes into contact with the liquid surface. Place typically infected larvae (10-30 as they fit) on the filter paper over the edge of the watch glass (Fig. 2). IJs will start to exit 10-12 days after infection. The goal is for active infectives to migrate into the water or formalin while most of the host tissues slough into the concavity of the watch glass. Once nematodes begin to appear, they should be harvested daily until production drops (3-4 days). To harvest, remove the watch glass with dead larvae, pour the IJs into a beaker (rinse the petri dish to collect all the lJs), add 70 ml of sdw or 0.1% formalin and replace watch glass.

For S. glaseri,'pre-IJs' are the form that exits from the cadaver. If these juveniles migrate into an entirely liquid medium, they will not successfully complete development to IJs. Good results have been obtained by replacing the watch glass and filter paper with a specially prepared petri dish. Prepare the petri dish by filling with a thin layer (2mm) of plaster of Paris, allowing the plaster to dry and then rewetting the plaster just prior to use. The plaster should be wet, but there should be no standing water. Monitor the moisture level throughout the harvesting period, and add a few drops of water as necessary. Infected hosts are placed on the wet plaster. Pre-lJs emerge, finish development on the plaster substrate and then migrate within 2-3 days into the water or formalin in the larger surrounding glass petri dish. They can then be harvested in the normal fashion.

3. Preparation for Storage: Examine the lJs for activity. Host tissues and noninfective stages should not be present. If extraneous matter or large numbers of inactive lJs are collected, a separatory step should be performed before the final rinsing procedures.

Separation of active lJs from other material is performed by allowing the IJs to migrate through a filter chosen to retain the host tissues and noninfective stages. Noninfective stages of steinernematids may first be killed by rinsing the nematodes in 0.4% Hyamine"I solution (Methylbenzethonium chloride) for 15 minutes. If desired, noninfective stages of both steinernematids and heterorhabditids may be killed by allowing the nematodes to sit at room temperature for a few days. In some cases, however, it will be simpler to use a fine mesh filter to retain the larger stages. Filters in use include 1) 3-4 stacked Kimwipesl" or a milk filter (FJeen Test Products Inc., Milwaukee, WI 53201) supported on a wire mesh in a Baermann funnel, 2) a 500-mesh (30-lim opening) screen in a separatory funnel, or 3) a 500-mesh sieve. In each case the filter is placed in contact with sdw in the collection container. Depending upon the numbers, most lJs should have migrated within 8 hours.

To rinse IJs, allow them to settle in a beaker. Then aspirate or decant the supernatant, and add more sdw until the suspension is clean (2-4 times). If the suspension appears particularly contaminated, it may be rinsed once with 0.1% formalin. Centrifugation at 300 rpm for 1 minute may be used to speed the settling process. When time is short, the nematodes may be stored overnight in a refrigerator but should then be rinsed at least one more time before storage. Finally, transfer the nematodes to a storage container.

In Vitro Production

In the past, steinernematids and heterorhabditids have been cultured on a variety of substrates: potato mash (McCoy and Glaser 1936), ground veal pulp (McCoy and Girth 1938) and dog food (House et al. 1965, Hara et al. 1981). Currently, a medium based on chicken offal (Bedding 1984) is common. The important factors seem to be monoxenicity (i.e., the nematode and associated bacterium as the only biotic agents), the use of primary form bacteria, a large surface area on which the nematodes may grow, a sterol source for the nematode and a food base for the bacterium. Further information on important aspects of in vitro rearing is available (Bedding 1986).

Bedding (1981, 1984) has developed a technique whereby huge numbers of nematodes may be economically produced using a chicken offal medium on a porous foam substrate. Polyether polyurethane provides the largest surface-to-volume ratio while providing adequate interstitial space (Bedding 1986). Glass flasks or large autoclavable bags serve as rearing containers. Proficiency with the flask system is considered a prerequisite to successful culturing in the bags, which average a production of over 1 billion S. feltiae IJs/bag (Bedding 1986). The method outlined below is for 500-ml widemouth Erlenmeyer flasks.

1. Preparation of Rearing Flasks: Wearing rubber gloves during this procedure is highly desirable. Impregnate small foam pieces (ca. 1-cm diameter) with chicken, duck or turkey off a homogenate (Appendix II, No. 3). Bedding (1984) recommends 12.5 parts medium to 1 part foam, by weight. The pores of the foam should still be clearly visible, but mediurr, should ooze out when the foam is squeezed. Fill the flasks with foam homogenate mixture to the 250- t( 300-ml mark (about 100 g). Wipe the mouth of the flasks well. Plug with cotton wrapped in cheesecloth, and autoclave for 20 minutes at 121 C. When the offal homogenate has been frozen prior to use, it can develop an offensive odor, so an autoclave deodorizer may be in order.

2. Inoculation with Bacteria: The day before the flasks will be prepared, liquid cultures of the primary form of the appropriate Xenorhabdus (Appendix 1. No. 1) should-be-incubated. Cells of the bacterium should be aseptically transferred to 5 ml of nutrient broth Appendix II, No. 1) in,a test tube (one tube per flask to be inoculated). If possible, leave the. tubes overnight on a shaker or at least vortex the tubes before incubating.

Allow the autoclaved flasks to cool to room temperature. Inoculate by pouring the contents of one tube of bacteria into each flask. Shake to mix the broth and bacteria throughout the foam substrate. Store for 2-3 days at 25 C to allow the Xenorhabdus population build up.

3. Inoculation with Nematodes: When monoxenic cultures are already available, foam from them is used to septically inoculate the new flasks. One flask can be divided into about seven new ones. Care should be taken to maintain mon oxenicity during th(transfers. Best results are obtained if the flask is not shaken after introduction of the nematodes. Th( flask will be ready to harvest in about 2 weeks. It is possible to employ a much smaller inoculum such aone or two pieces of foam. However, harvest will then be delayed to 4 weeks, and the final yield will be reduced. During the extra 2 weeks, the proportion of secondary Xenorhabdus and concentration of metabolites increase so that the final population is less than for those flasks with a higher initial inoculation rate.

When monoxenic cultures are not available, or when their purity is suspect, surface-sterilized infectives (Appendix 1, No 6) may serve as the inoculum. Do not..add-too.much liquid with the nematodes (< 5ml). When beginning with newly sterilized IJs, the purity of the new cultures should be verified within 2-3 ' days. Aseptically streaking from the flask onto MacConkey agar or NBTA (Nutrierit agar, bromothymol blue and tetrazolium chloride (Appendix 11, No. 1)) and then checking for appropriate colony color (Appendix I, No. 1) and morphology should generally be sufficient to verify purity (Table 5). Cell morphology (rod-shaped, see Table 5 for dimensions) and motility may also be examined if desired.

4. Harvesting: The foam may be piled 5 cm deep on a 20-mesh sieve (20 meshes/inch). Place sieve in a pan of tap water with water level adjusted so that the foam is just submerged. If a mist chamber is available, the pan plus sieve may be placed in it for 2-24 hours. (Longer periods tend to be detrimental to the nematodes.) Do not pour water over the foam as this washes particles of homogenate into the water. Within 2 hours 95% of the lJs will migrate into the water (Bedding 1984).

The nematodes may be sedimented and rinsed if necessary to remove particulate matter and inactive lJs. The lJs may then be permitted to migrate through a 500-mesh sieve. Nematodes rinsed from the inside of the flask should also be allowed to migrate through the 500-mesh sieve to remove particulate matter. (For steinernematids, the non-ljs may be killed in a Hyaminel" solution before migration through the sieve.) The nematodes should be rinsed several times until the water appears clear. An antibiotic rinse may be deemed appropriate (see part 3 of "In Vivo Production" above). Rather than rinsing several times right after harvesting, Bedding (1984) first aerated steinernematids for 1 week to break down small particles of medium then rinsed the lJs.

5. Troubleshooting: A drop in production could indicate contamination, a reversion to the secondary phase of Xenorhabdus, unsuitable incubation temperatures, improper moisture content or many other problems.

Contamination ivill sometimes be visually evident in'the form of fungal or bacterial colonies or'unusual' coloration or exudates. At other times an 'unusual' odor may indicate that contamination is a problem. In any case, purity should be routinely verified by streaking onto NBTA or MacConkey agar (Appendix i, No. 1). Using media with specific dyes also permits determination of the proportions of primaryand secondary-form bacteria. By the end of 2 weeks, it is normal for a flask to contain a high proportion of secondary bacteria; however, it is important that the original bacterial inoculation for each flask be done with primary-phase bacteria.

As to incubation temperatures, 25 C seems optimal for S. feltiae, but the optimum may be different for different species. Room temperatures of 20-25 C should generally be adequate for good production of nematodes.

Moisture content can sometimes be difficult to adjust. It -may be necessary to try adding more or less liquid at various steps in the procedure. If the medium is too liquid, or if too much liquid is added, the nematodes will become trapped in standing water and die. If the medium is too dry, the cultures will dry out before large numbers of nematodes are produced. Maintaining high humidity where flasks are incubated may be helpful.

Production levels can remain adequate while quality, measured in terms of infectivity or activity, drops. In vivo passage is recommended every 4-6 months to avoid reduced virulence and pathogenicity of the nematodes to their insect hosts. Otherwise, strict attention to maintaining optimum conditions for growth, development and storage will reduce the potential for problems with nematode quality.

It is impossible to foresee every potential problem. However, each technician will become familiar with his/her own system and its responses to procedural changes. With some knowledge of nematode biology, logical causes and solutions for most troubles should not be too difficult to find. Even so, in uitro production is not easy, and trial and error will be needed for the novice to perfect the system.

6. Primary-Phase Xenorhabdus Cultures: Primary- phase bacteria may be isolated and recognized by methods in Appendix I, No. 1. Primary colonies may be suspended in 17% glycerounutrient broth in Martney bottles and stored at - 18 C (Akhurst 1980). In this case the suspension is rapidly thawed in a water bath at 60 C prior to use (Bedding 1984). Alternately, the colonies can be stored at 12-25 C and routinely subcultured (monthly at 12 C or weekly at 25 C). It is best to use NBTA or MacConkey agar so that colonies reverting to the secondary form will be easily recognized and will not be used.


The nematodes may be stored in distilled water with a droi) of Triton X-100 (a wetting agent that prevents nematodes from sticking to the side of the container) or 0.1% formalin (use this only if continuing or recurring contamination seems to be a problem). If stored without aeration, the nematodes should be concentrated to no more than 10,00020,000 nematodes/ml, and the water depth should be 1 cm or less. Tissue culture flasks are ideal for this type of storage since there is a large surface-tovolume ratio.

Higher nematode concentrations (100,000/ml) will not be detrimental in aerated suspension. An aquarium pump or any forced air supply can be attached to an aquarium stone (a porous structure that creates many fine bubbles as the air is forced through) in the nematode suspension.

Steinernematids can be stored at 4-10 C for 6-12 months without much loss of activity. Heterorhabditids do not store as well, and 2-4 months of storage at 4-10 C is considered good. Pye and Burman (1981b) report the formation of rosettes (clumps) of heterorhabditids. Addition of sodium bicarbonate solution (stock solution of 1 g NaH,CO., per 50 ml H,O, or stronger) breaks up the rosettes without any other apparent effect on the nematodes (Timper unpub. obsv.).

Bedding (1984, 1986) also proposes the following storage procedure. For up to 1 year, a well-rinsed semisolid paste of steinernematid lJs may be stored on moist autoclaved polyether polyurethane foam. Coat the foam with 10 times its weight of IJs. Place the foam in a sterilized container and force air through bacterial filters (0.45 microns). The antiseptic precautions are important to minimize fungal and bacterial contamination. The lJs can be extracted later by migration through a sieve as during the harvesting process. Or, foam may be squeezed in water until all the tan 'nematode' coloration is gone. Extraction is overly time consuming for largescale commercial use but poses few problems for research purposes. S. feltiae lJs are extracted most quickly at 28 C (Kirkpatrick pers. comm.).

Storage in activated charcoal (Yukawa and Pitt 1985) or in an inactive dried state are other new possibilities.

Transport of Live Nematodes

Live IJs may be shipped as a semisolid paste on moist foam (as they are stored) or cotton. The container should be kept cool (packed in ice, not dry ice), and shipment must be expedited to enhance survival. The use of overnight delivery systems is encouraged. Shipment in water is no longer recommended because better aeration is attained on the moist foam or cotton.