General Considerations for Determining the Epidemiology and Control of Gastrointestinal Nematodes in Wild and Exotic Ruminants
Department of Veterinary Pathobiology, Texas A&M University, College Station, TX, USA
Despite the seemingly endless names of gastrointestinal parasites in various species of wild and exotic ruminants the parasites can generally be divided into two categories. These categories are based on seasonal transmission patterns, either cool season or warm season. Depending on the weather conditions in a given geographic area, the seasonality of transmission may be prolonged or short to nonexistent for a particular parasite species. Each parasite species has optimal conditions for development from egg to infective larvae. Not only are there optimal conditions but there are temperature and moisture limits for development and subsequent survival in pastures.
The vegetation that grows best in an environment is usually cool or warm season forages and the transmission of parasitic nematodes can be associated with different vegetation growth. Haemonchus spp., a warm season parasite, is associated with bermudagrass and growth in the summer. Ostertagia spp., a cool season parasite, with rye grass growth in the autumn and spring. When these forages are dormant, the larvae either quickly die or become inactive until conditions in the pasture become favorable for transmission.
Studies done in Texas utilizing tracer lambs were valuable in determining the transmission of gastrointestinal nematodes in free ranging exotic ruminants. The species of gastrointestinal nematodes found in antelope readily infect domestic sheep. Lambs, free of internal parasites, were sequentially grazed, for approximately 1 month, in pastures with antelope. They were then slaughtered and the species of worms and developmental stage present were determined.
The tracer lambs acquired worms in a pattern similar to that of domesticated sheep in the same region.1 The primary warm season parasite was Haemonchus contortus. In the southern United States this parasite can be transmitted during any month of the year provided that there has been greater than 50 mm of precipitation and a daily mean temperature above 10°C. However, the greatest level of transmission is between April and November with the peak varying with the rainfall pattern of the year. There seemed to be a constant adult worm population and arrested development was not identified under the experimental conditions but may be important in some situations. Cold weather conditions do not kill the free-living stages but may slow development sufficiently that the egg or first and second larval stages desiccate in the environment. Infective larvae can over-winter on pasture, even if inactive, until the weather warms up and transmission occurs. Haemonchus larvae survive the cool season in pasture for 6 months or longer but will die in 1–2 months in the summer as the energy sources are depleted.
The primary cool season parasites of antelope are Ostertaginae (Camelostrongylus and Longistrongylus) and Trichostrongylus spp. which are primarily transmitted from September to May. Peak transmission occurs in October and November and again in March, April, and May. The Ostertaginae of cervids (Spiculopteragia, Ostertagia) also appear to follow this pattern. Cool season and warm season parasites overlap at times and as one goes farther north, the more the transmission patterns become similar. Cool season parasites are not cold season and are inactive during the winter but quickly become active in the spring. They do not survive hot dry summers in pasture. Arrested development of Camelostrongylus mentulalus was seen in tracer lambs. In Texas this was a summer arrest that differs from a winter arrest for this species reported in a Scottish zoo.2
When an animal dies, for whatever reason, one of the most important findings on postmortem is to determine which species of parasites are present and the abundance of each. When you know which worms are laying, those eggs that look the same, and the general patterns of transmission, then you might be able to select a rational approach to helminth control. The off-season, when parasites are in the host, is the important time to treat, not when larvae are being transmitted in the greatest numbers. During the off-season treatment with an effective anthelmintic may remove 10–40% of the total worms in the environment whereas treatment during the peak of transmission will only affect 0.1–1% of the total worm population. You may remove thousands but leave billions on the pasture.
Treatment with traditional anthelmintics may remove adult and/or immature worms within the host at that time. Treatment with an anthelmintic with residual activity should, in theory, remove not only the worms in the body but also those acquired from the environment for the next few weeks to 1 month. When administered during the off season it will perhaps do this. However, injectable anthelmintics with residual activity can also be a potent mechanism for selection of drug resistant nematodes. As the anthelmintic blood level falls those worms with a semi resistant genotype will establish in the host and mate with similar worms selecting for resistant worm populations.
Anthelmintic resistance is a problem in goats, exotic wildlife, and sheep in that order. Where the pharmacodynamics of an anthelmintic is known, goats and at least some exotics (e.g., red deer) require a higher dose to deliver the same level of exposure by the drug to the worm than in sheep or cattle. Anthelmintics used in exotic populations may have sufficient activity to produce a clinical response but the effectiveness will fall with each worm generation until the drug becomes useless. Clinical response or fecal egg counts before and after treatment has been the traditional method of anthelmintic evaluation. Recently a combination egg hatch and larval development assay was developed which is able to predict the susceptibility of worm populations to anthelmintics. Anthelmintics cause deleterious effects in the free-living developing stages similar to that of parasitic adult worms.3 Nematode eggs are incubated in solutions of varying concentrations of anthelmintic, when a concentration is reached where the egg is either unable to hatch or to prevent larvae from developing to the infective stage, the adult worms of this population are similarly affected. The test does not evaluate variations in the host’s metabolizing of the drug but indicates the susceptibly of the worm to the drug. If the eggs in control wells can develop to infective larvae in 1 week in 100% humidity at 26°C the species of parasites susceptible to the drug can be identified. The test indicates which genera of worms are affected by anthelmintics so that important target species can be controlled. The determination of which anthelmintics (benzimidazoles, levamisole, macrocyclic lactones, and combinations) are likely to be effective are evaluated in a single test. The larvae development assay can tell you if the problem is:
1. The wrong anthelmintic, that no matter how given would not work
2. The drug is fine but the dose, method of administration, or other host factors is the problem
The key to controlling gastrointestinal nematodes in exotics is:
1. To know the parasites you are dealing with
2. To know when the parasites are being transmitted
3. Treat animals in the off season
4. Use effective anthelmintics
5. Giving an adequate does in an effective way to the host species
We also must remember that the parasites are largely shared and some hosts are little affected by certain species of parasites which are devastating in others. All of the hosts in the ecosystem (ranch, park, enclosure, pen) must be evaluated, not just the rare expensive individuals.
1. Craig TM. Epidemiology and control of gastrointestinal nematodes and cestodes in small ruminants: southern United States. Vet Clin North Am Food Anim Pract. 1986;2:367–372.
2. Flach EJ, Sewell MMH. Gastrointestinal nematodiasis in black buck (Antelope cervicapra) at Edinburgh Zoo. J Zoo Anim Med. 1987;18:56–61.
3. Maingi N, Bjorn H, Dangolla A. The relationship between fecal egg count reduction and the lethal dose 50% in the egg hatch assay and larval development assay. Vet Parasitol. 1998;77:133–145.