Epidemiology of GI Parasites in Ruminants
All grazing animals become infected with GI nematodes during their lifetime. The severity of the resulting disease can range from no clinical signs to debilitating, fatal disease, depending on the strength of the animals’ acquired immunity, the magnitude of exposure, and concurrent stressors or illness.
Ruminants acquire protective, non-sterilizing immunity to GI nematodes over time, after exposure to the parasites. Therefore, young, growing animals are the most susceptible to disease, especially in the first pasture season. Although the presence of nematode-specific antibodies has been shown in the milk and colostrum of cattle and small ruminants, it is not known whether these antibodies induce protective immunity. Animals kept in an environment where they are not exposed to parasites do not develop protective immunity, and they suffer severe clinical signs, even as adults, on first exposure.
In addition, it has been well established that a small proportion of the animals in a herd shed the majority of eggs (overdispersion). These prolific parasite shedders, which are a large source of environmental contamination to the rest of a herd, can be identified and managed, or culled accordingly.
Young ruminants also acquire infections caused by cestode parasites (Moniezia, Thysanosoma) while grazing. Cestodes have an indirect life cycle that requires the common orbatid mite as an intermediate host. Mild to moderate infections with cestodes rarely produce clinical signs, and acquired non-sterilizing immunity generally prevents disease, unless the animal carries very heavy burdens.
Other factors that affect the presence of clinical signs of GI parasitism include stressors or illnesses that may impair the immune system response. Examples include late pregnancy, lactation, weaning, shearing, overcrowding, inadequate nutrition, and primary illnesses. A periparturient rise in egg counts occurs for most GI parasites during late pregnancy and early lactation. This periparturient rise can represent an additional source of infection for neonates, or increased pasture contamination for herdmates.
Beginning in the early 2000's, the concept of "preserving the refugia" in parasite management was introduced. A refugium represents the proportion of parasites in a population that have not been exposed to a dewormer. Parasites in refugia that have escaped deworming can reproduce and contribute alleles to the next generation of parasites on the pasture. With large refugia, susceptible alleles (worms susceptible to dewormer) dilute resistance alleles, and resistant parasites are less likely to reproduce with other resistant worms. Refugia-based strategies will slow resistance evolution in a herd. Sources of refugia include parasite eggs and larvae on pasture or living in untreated animals.
A final key point is that some GI parasite larvae (such as Ostertagia L3 and L4) can enter an "inhibited" stage in either the environment or the host, also referred to as "hypobiosis". This period of arrested development can complicate the interpretation of fecal egg counts, as they will not represent the true (potential) parasite burden in the animal. As pertains to the environmental management, pastures considered to be "clean" of parasites after winter may still be contaminated with infective organisms.
Features and Classification of GI Parasites in Ruminants
Most parasites that infect the GI tract of ruminants are nematodes; occasionally, however, some pathogenic cestodes—eg, Moniezia and Thysanosoma—are encountered. Coccidia of ruminants, as well as other GI protozoa, such as Giardia and Cryptosporidium, are discussed elsewhere (see Giardiasis and Cryptosporidiosis).
Because most GI parasites are nematodes, their life cycles, clinical signs, and control programs are similar. All grazing animals will become infected with GI parasites during their lifetime, but infection does not always correlate with disease or production losses.
Efforts should be made to identify populations of parasites present within individual herds because parasite prevalence and magnitude of infection can vary by geographic region, local climate, individual management factors, and historical deworming practices. After patterns of parasites in a herd have been identified by means of integrative diagnostics, a control program should be implemented to prevent disease and noteworthy production losses.
For practical purposes, GI nematodes of ruminants can be divided into two major categories:
Trichostrongyle parasites (order Strongylida, also known as strongyles) include numerous genera, such as Ostertagia, Teladorsagia, Haemonchus, Trichostrongylus, Cooperia, Nematodirus, Mecistocirrus, Bunostomum, Oesophagostomum, and Chabertia. At the typical diagnostic stage, these trichostrongyle parasites (eggs on fecal flotation) are indistinguishable. Their life cycles are generally the same, differing only in slight nuances, such as the length of each stage, their location in the GI tract of adults, or their preferred environmental temperature ranges. Life cycles of all the trichostrongyles are direct and rely on animals to graze contaminated pastures to initiate infection. For these reasons, their diagnosis, treatment, control and prevention are very similar.
Nontrichostrongyle parasites include Strongyloides, Trichuris, Aonchotheca, and Toxocara vitulorum. Each of these nontrichostrongyles has a unique life cycle, exhibits distinctive eggs on fecal flotation, and may have different treatment, control, and prevention methods than trichostrongyle parasites have.
Courtesy of Dr. Grace VanHoy.
The generic life cycle of a trichostrongyle begins when an adult female in the host GI tract mates with an adult male and produces fertilized eggs. Females of each trichostrongyle species have differing reproductive capacity; eg, Haemonchus contortus females have been shown to produce up to 10,000 eggs per day, and Nematodirus spp females produce < 100 eggs per day. The wide variation in fecundity of different trichostrongyle species affects the utility and reliability of diagnostic tests.
Eggs are passed in the feces of the host into the environment and continue their development in the fecal pat or pellet. Under ideal conditions—between 10°C and 36°C (50°F and 96°F)—eggs hatch into first-stage larvae (L1s) in 1–2 days. Conventional belief has been that climates with temperatures outside of this range allow no substantial accumulation of parasites on pasture, although recent evidence suggests otherwise.A notable exception is Nematodirus, which overwinters in the soil and requires a period of cold to develop. Unlike other trichostrongyles of ruminants, Nematodirus larvae develop within the egg, and animals are infected when they ingest the hatched larvae.
In addition to ideal temperatures, moisture is critical for larval development. Humid, temperate climates favor the rapid development of trichostrongyles; however, arid climates can also support parasite development if irrigation is practiced. After eggs hatch, the L1 continues developing in the feces and molts into second- and third-stage larvae (L2, L3) over 4–6 days. In ~7 days, infective L3 larvae migrate from the feces and begin to climb vegetation, where they are consumed by the host. Once inside the host, L3s complete two more molts to reach adulthood inside the abomasum or small intestine.
The prepatent period for most trichostrongyles is between 18 and 21 days, meaning that eggs can be detected in host feces from a new infection 18–21 days after ingestion of the L3. Unique life cycle adaptations for the abomasal parasites (Ostertagia and, to a lesser extent, Haemonchus) help these species overcome poor environmental conditions. If the weather is cold, L4s invade the gastric pits in the abomasum, and development is arrested. Once the weather is favorable again, L4s undergo a massive, coordinated emergence that leads to severe disease of the host. The resulting syndrome is called type II ostertagiasis, as discussed in Clinical Findings.
Several life cycle nuances can be manipulated by owners or veterinarians to predict high-risk periods for their animals or change management factors to decrease the risk of heavy infection. When in the environment, L3 larvae have a tough outer membrane called a cuticle, which protects them but also prevents them from feeding. If an L3 does not feed before its finite energy sources run out, it will die. L3 metabolic rates change according to environmental conditions: cool, dry weather decreases the metabolic rate and prolongs the L3 lifespan; hot weather increases metabolic rate and decreases lifespan.
Recommendations for pasture rest times can be based on these factors; pastures can be rested for shorter periods of time when it is hot, but they need to be rested for longer periods of time in cool, dry weather. Similarly, humid, mild temperatures and rainfall events in temperate-arid climates increase the success of trichostrongyles and accelerate the rate of their development on pasture. Increased monitoring is required during these high-risk periods. In addition, trichostrongyles require vegetation to complete their life cycle, so animals on a dry lot will have a substantially decreased risk of detrimental infection.
The patterns and prevalence of specific GI parasites are regionally and climactically specific. However, differing patterns have emerged because of the shipping of animals over long distances, the effects of global climate change on parasite life cycles, and the susceptibility of parasites to certain anthelmintics.
Clinical Findings of GI Parasites in Ruminants
Most GI parasite infections in ruminants are mixed, with several genera infecting a single animal. Notable exceptions are haemonchosis and type II ostertagiasis . In general, clinical signs are present in only a small proportion of a group of animals, even though all the animals have GI parasites. The clinical signs associated with GI parasitism are typically nonspecific and manifest as a chronic, slowly progressive illness unless a very high infectious dose is encountered or other concurrent risk factors are present.
Heavily infected animals experience gradual weight loss or decreased weight gain (in the case of young animals), ill thrift, gradually decreasing milk production, and poor hair coat. More severely affected animals show lethargy and weakness, and they may spend more time away from the group or lying down. Emaciation may occur in long-standing cases because of inappetence that progresses to anorexia, as well as poor nutrient absorption. Chronic GI parasitism may also predispose animals to secondary infections (bacterial, viral, fungal) that they would normally be able to resist.
Diarrhea may be present, depending on which segment of the GI tract is most heavily parasitized. Ruminants with predominant Haemonchus infection of the abomasum may have formed feces until late in disease; ruminants with predominant Trichuris infection of the colon will have diarrhea early in disease. Occasionally, some ruminants display signs of colic (abdominal discomfort, kicking at the abdomen, rolling, repeated bouts of standing up and lying down).
Whether or not the animal has diarrhea, heavy GI parasitism causes a protein-losing enteropathy as a result of either increased loss from diarrhea or decreased absorption from direct damage to the mucosa by the parasites. Clinical signs of hypoproteinemia might include submandibular edema (bottle jaw) or ventral edema. In addition to inadequate protein absorption, heavily parasitized ruminants are likely to show poor absorption of critical vitamins and minerals from their diet.
Bloodsucking species of trichostrongyles (eg, Haemonchus spp) cause severe anemia; however, some anemia can also be found with most heavy trichostrongyle infections. Because trichostrongyles use cutting or slicing mouthparts to attach to abomasal or intestinal mucosa, blood can leak from these sites. Clinical anemia is commonly chronic, and many animals can compensate until their hematocrit is < 10%. Severe anemia presents as cardiovascular collapse: increased heart rate, increased respiratory rate and effort (open-mouth breathing, cheek puffing), poor perfusion, cold extremities, and sudden death. Acute decompensation may follow a quicker clinical course after heavy exposure (eg, after large rainfall events that accelerate larval development), or after an animal with chronic parasitism reaches critical levels of anemia, hypoproteinemia, and/or emaciation.
Diagnosis of GI Parasites in Ruminants
Infection with GI parasites can be confirmed by demonstration of nematode eggs or tapeworm segments on fecal examination. Fecal examination can be further separated into qualitative and quantitative evaluations (see below). Feces should be fresh, and samples are ideally collected before they come in contact with the ground, to avoid environmental contamination with free-living stages of nematodes. If an individual animal is suspected of disease, feces should be collected directly from the rectum.
Qualitative Fecal Evaluation
A centrifugation fecal flotation performed on individual animal feces or on composite samples from multiple animals within a herd enables a qualitative assessment of the types of GI nematodes shedding eggs in the feces and a subjective assessment of the magnitude of infection. Composite fecal sampling is a robust method for evaluating parasite burden within a herd that is a resource-efficient option for owners and producers.
To avoid development of the parasite eggs, fecal samples must be stored in an airtight container in a refrigerator and used within 5 days. Several flotation solutions are validated for use with ruminant feces. If parasite eggs have a specific gravity less than the specific gravity of the flotation solution, they rise to the top. These recovered parasites are transferred to a glass slide and examined by microscopy. False negatives can occur if an egg is more dense than the solution chosen, and thus does not float.
Several commercially available sugar and salt flotation solutions are validated for use in ruminants. Each flotation solution has a different specific gravity (and therefore different recovery of specific parasites) and a different shelf life.
Sugar solutions do not distort egg morphology, and they can be examined after several hours or days, if refrigerated; however, these solutions have a shorter shelf life than salt solutions have, and they can be prone to fungal or bacterial growth unless a preservative is added. Salt solutions have a longer shelf life but must be examined shortly after preparation because slides will crystallize, disrupting or destroying parasite egg membranes.
Most common GI nematode eggs will be recovered if a flotation solution has a specific gravity of at least 1.24 (Sheather’s sugar flotation solution has a specific gravity of 1.26–1.27). Passive flotation is no longer an acceptable standard of practice.
Quantitative Fecal Evaluation
Several fecal egg-counting techniques have been validated for use in ruminants and have different levels of sensitivity. Examples of techniques include modified Stoll’s, modified McMaster’s, Wisconsin, FEKPAK (Australia), and mini-FLOTAC. Fecal egg counts are a more objective measure of the number of parasite eggs than is fecal flotation, and they can indicate the efficacy of dewormers (discussed in Treatment). Fecal egg counts can provide robust information about herd health, and with good record keeping they can create a strong clinical picture of infection patterns within herds over time. Veterinarians and owners can use these patterns to establish treatment thresholds.
Factors that affect the number of eggs in a sample include the relative fecundity of female worms, which varies with the nematode genus; the existence of immature nematode stages not yet producing eggs but still creating damage to mucous membranes within the GI tract; diarrhea (dilution of eggs); and the recent use of dewormers. In addition, some egg-counting techniques require specialized equipment and fail to identify the genus or species of trichostrongyle-type eggs. For these reasons, fecal egg counts should not be the sole determinant of treatment and management of clinical GI parasitism.
Genus- and Species-Level Determinations
Specialized techniques can definitively determine the genus and species of the GI nematode infecting individual animals or a herd. This information may be helpful in initially evaluating herd-level parasitism, or when new problems are encountered, but these techniques are rarely practical.
Larval culture of the feces (coproculture) enables the eggs to develop to infective third-stage larvae (L3s), which can be identified by microscopy. Fecal flotation slides can be labeled with a fluorescein peanut agglutinin and observed for fluorescence (only Haemonchus eggs fluoresce). Fecal samples can also be evaluated by PCR assay to determine the specific nematode genus and species. All of these techniques are generally performed by a specialized laboratory.
Postmortem Examination
All animals suspected to have died as a result of GI parasitism should undergo a postmortem examination. Each section of the GI tract should be examined for mucosal defects, hemorrhages, erosions or ulcers, and the presence of nematodes or cestodes attached to the mucosa. These parasites can be very small and threadlike, and each species has a preference for a particular GI tract section.
GI contents can be collected and run through screens to identify parasite larvae or adults. Larvae and adults can be identified to the level of genus with microscopy by specialty labs; molecular techniques make identification of the species possible. Histopathology should be conducted on abnormal sections of the GI tract to identify parasites embedded in the mucosal wall.
Ancillary Data
Major bloodwork abnormalities in ruminants clinically affected by GI parasites include anemia and hypoproteinemia characterized by hypoalbuminemia. Eosinophilia on the complete blood count differential may be present; however, a normal result cannot rule out clinical parasitism. Animals with severe, acute infection with diarrhea may have associated electrolyte abnormalities, including hyponatremia, hypochloremia, hypokalemia, and metabolic acidosis characterized by bicarbonate loss.
In animals with abomasal parasitism, abomasal pH may increase as acid production decreases secondary to parietal cell damage. Samples may be obtained by abomasocentesis. Pepsinogen, the inactive form of pepsin, is not activated if the abomasum is not acidic. Increased concentrations of serum pepsinogen may be found in cases of severe type II ostertagiasis; however, few diagnostic laboratories in the US offer this test anymore.
Integrated Parasite Management
"Integrated parasite management" involves a strategic and often multi-faceted approach, and plans vary with herd and farm. A comprehensive knowledge of the parasites infecting the herd, the parasite biology, and risk factors present in the environment and in the animals themselves should all be taken into account. Integrated parasite management plans include the following components:
Prevention
Monitoring
Treatment
Refugia
Biosecurity and quarantine
Alternative (nondrug) adjunctive options
Prevention of GI Parasites in Ruminants
Infection with GI parasites does not equate to disease; in fact, animals that are not exposed to a low load of GI parasites will never form protective immunity and are more at risk of severe disease when exposed. For this reason, preventive measures should center on preventing disease rather than eliminating all parasites.
All factors in the triad of epidemiology (host, environment, pathogen) can be addressed in a prevention program. Host factors that aid in the avoidance of clinical GI parasitism include adequate nutrition (especially protein); adequate vitamin and mineral supplementation, if needed to support the immune system; prevention of common diseases that result in immunosuppression; and prevention of stress. Addressing host factors might include paying special attention to bunk space, overcrowding, mixing of social groups, and common stressors such as weaning, vaccination, shearing, etc.
Resistance to GI parasitism and immune response to GI parasite challenges are heritable traits that have been studied extensively in sheep. Recommended methods include breeding animals that seem resistant to GI parasite burdens, and culling poor doers from a breeding program.
The free-living stages of GI parasites on pasture, and the effect of environmental management on parasite epidemiology, make pasture management a critical tool for effective prevention of disease. Susceptible animals (young animals without acquired immunity, pregnant animals, sick animals, etc) should not be exposed to pastures that are potentially highly contaminated with parasites. Exploiting the typical life cycle and characteristics of GI nematodes can aid in effective pasture management. Here are some examples:
Trichostrongyles speed up their life cycle and increase their population under ideal environmental conditions.
Intervention: Remove grazing animals from pasture after rain events, or during humid and mild conditions.
Infectious L3s can vertically migrate only short distances and are found close to the ground.
Intervention: Graze only pastures with forage longer than 10 cm.
Production animals do not share pathogenic GI nematodes with other nonruminant species and can help clear pastures of parasites for each other.
Intervention: Co-graze or alternately graze high-risk pastures with different species (eg, horses).
Drug-resistant parasites survive treatment with certain drugs and may be the only population left after deworming, contaminating a “clean” pasture.
Intervention: Avoid deworming the entire herd at one time, and do not move the animals to clean pasture immediately after deworming.
Trichostrongyles require pasture and cannot complete their life cycle when exposed to high heat, desiccation, or prolonged direct sunlight.
Intervention: House animals on concrete or dry lots during high-risk periods or in cases of severe multidrug resistance.
Trichostrongyle L3s have a finite lifespan that depends on environmental conditions.
Intervention: Allow pastures to rest for 3–6 months (depending on the climate) between grazings.
Monitoring of GI Parasites in Ruminants
All ruminants maintain a burden of GI parasites during their lifetime, but infection is not equivalent to disease. To avoid disease, animals with increasing burdens must be detected. The amount of recommended monitoring depends on the herd, because production metrics and management are specific to each; ideally, however, monitoring should occur multiple times each year and particularly after stressors such as shearing, vaccination, weaning, and weather events that may favor the development of infective larvae on pasture.
Body condition scores and production metrics such as average daily weight gain, milk volumes, and activity levels can be monitored at low cost, and they are valuable information if good records can be kept. Fecal egg counts (individual or composite) and fecal egg count reduction tests (discussed in Treatment) can be an important part of a monitoring program; however, they require more resources and represent additional cost. Because ELISA can be used to detect Ostertagia antibodies in bulk-tank milk samples on dairies, it may also be a method by which herds can be monitored to determine infection prevalence and inform strategic deworming decisions.
Courtesy of Dr. Grace VanHoy.
Evaluation and tracking of FAMACHA scores (ratings of signs of pallor on the ocular mucous membranes that indicate anemia) can be a highly effective tool in determining when to deworm individuals in a herd affected with parasites that cause anemia, especially Haemonchus. FAMACHA scoring correlates the color of ocular mucous membranes to a numerical scale. Examiners compare the eyelid conjunctiva by covering the eye with eyelids, gently retropulsing the globe, and then pulling the lower eyelid conjunctiva and holding it next to a scorecard. Higher scores reflect paler mucous membranes, indicating anemia. Owners and veterinarians can be trained to perform FAMACHA scoring and obtain a scorecard easily with online training administered through the University of Rhode Island and the American Consortium for Small Ruminant Parasite Control (ACSRPC).
Although GI parasitism is not the only cause of anemia, the FAMACHA scoring system is highly correlated with the prediction of anemia from the bloodsucking trichostrongyle Haemonchus contortus in small ruminants and camelids. Informed, impromptu treatment decisions can be based on a combination of FAMACHA scoring, relative risk, and clinical examination without a reliance on fecal diagnostics. In addition, FAMACHA scores and other metrics, such as body condition score, can be recorded for each animal and monitored over time to detect seasonal trends in a specific herd.
The use of FAMACHA scoring has been found to markedly decrease dewormer use, increase production factors, and slow the development of anthelmintic resistance.(1)
Treatment of GI Parasites in Ruminants
Historically, owners and producers administered anthelmintics (dewormers) as a primary means of controlling GI parasites in their herds; however, severe and widespread multidrug resistance in the US and other countries has resulted in treatment failures and increased mortality. Because drug resistance evolution will outpace the development of new drugs, the efficacy of existing drugs should be carefully protected.
Although drugs have been the main historical component of GI parasite control, overuse and subsequent resistance have forced a paradigm shift in parasite management. Treatment strategies should be modified and anthelmintics administered more judiciously within an integrated parasite control program that is individualized for each herd. Anthelmintics should be administered only to treat clinically affected animals and decrease pasture contamination, especially at key points in the parasites’ life cycle. Monitoring tools (discussed above) can be used to identify both the clinically affected animals and the parasite(s) involved so that the specific parasites' biology is taken into account. All efforts should be made to maintain large refugia.
Treatment of Gastrointestinal Cestodes
Treatment of Moniezia and Thysanosoma is rarely warranted, given that tapeworms are generally nonpathogenic and ruminants develop a strong acquired immunity after the age of 5–6 months. In the US, fenbendazole, oxfendazole, and albendazole are approved for the treatment of cestode infestations in cattle, with some age restrictions.
Treatment of Gastrointestinal Nematodes
When treatment of parasitic infestation in ruminants is necessary, several options exist. In the US, cattle can be treated with a wide range of drug classes and formulations; goats and sheep, however, are considered a minor-use species, and the list of choices is narrower. All treatment decisions should be supported by the Food Animal Residue Avoidance Databank (FARAD) and the 1994 Animal Medicinal Drug Use Clarification Act (AMDUCA).
Anthelmintic administration may precipitate death in very sick animals, and owners should be warned of this potential complication. Careful dosing of anthelmintics should be based on accurate weights for each individual animal because underdosing can rapidly lead to anthelmintic resistance, and overdosing may have health consequences for the animal or economic costs for the owner.
In the US, resistance of GI nematodes to every drug class has been documented, especially in the Southeast. For this reason, anthelmintic efficacy should be monitored carefully with posttreatment diagnostics and confirmation that clinical signs have resolved. Rotational deworming is no longer recommended, because it has been documented to perpetuate the evolution of multidrug resistance.(2, 3) Combination deworming (administering two or three drugs from different drug classes simultaneously) can be used on farms with documented resistance. When evidence indicates that resistance to a particular drug exists on a farm, other drugs in the same drug class should be avoided because they share the same mechanism of action.
Drugs for the Treatment of GI Parasites in Cattle
--------------------GI Tract Location------------------- | |||
|---|---|---|---|
Drug Class | Abomasum (Nematodes) | Intestine (Nematodes) | Intestine (Cestodes) |
Benzimidazoles | |||
Albendazole | Yes | Yes | Yes |
Fenbendazole | Yes | Yes | No |
Oxfendazole | Yes | Yes | Yes |
Thiabendazole | Yes | Yes | No |
Imidazothiazole | |||
Levamisole | Yes | Yes | No |
Tetrahydropyrimidines | |||
Morantel | Yes | Yes | No |
Pyrantel | No approvals | ||
Macrocyclic Lactones | |||
Ivermectin | Yes | Yes | No |
Doramectin | Yes | Yes | No |
Moxidectin | Yes | Yes | No |
Eprinomectin | Yes | Yes | No |
Salicylanilide | |||
Closantel | No approvals | ||
Organophosphate | |||
Haloxon | Yes | Yes | No |
This table is based on information from the Food Animal Residue Avoidance Databank (FARAD), obtained May 8, 2023. Label claims regarding specific parasite species or genera as well as susceptible life stages (adult vs. larval vs. inhibited stages) must be observed. Resistance has been reported in all drug classes listed. | |||
Drugs for the Treatment of GI Parasites in Sheep
--------------------GI Tract Location------------------- | |||
|---|---|---|---|
Drug Class | Abomasum (Nematodes) | Intestine (Nematodes) | Intestine (Cestodes) |
Benzimidazoles | |||
Albendazole | Yes | Yes | Yes |
Fenbendazole | No approvals | ||
Oxfendazole | No approvals | ||
Thiabendazole | Yes | Yes | No |
Imidazothiazole | |||
Levamisole | Yes | Yes | No |
Tetrahydropyrimidines | |||
Morantel | No approvals | ||
Pyrantel | No approvals | ||
Macrocyclic Lactones | |||
Ivermectin | Yes | Yes | No |
Doramectin | No approvals | ||
Moxidectin | Yes | Yes | No |
Eprinomectin | No approvals | ||
Salicylanilide | |||
Closantel | No approvals | ||
Organophosphate | |||
Haloxon | No approvals | ||
This table is based on information from the Food Animal Residue Avoidance Databank (FARAD) obtained May 8, 2023. Label claims regarding specific parasite species or genera as well as susceptible life stages (adult vs. larval vs. inhibited stages) must be observed. Resistance has been reported in all drug classes listed. | |||
Drugs for the Treatment of GI Parasites in Goats
--------------------GI Tract Location------------------- | |||
|---|---|---|---|
Drug Class | Abomasum (Nematodes) | Intestine (Nematodes) | Intestine (Cestodes) |
Benzimidazoles | |||
Albendazole | Yes | No | No |
Fenbendazole | Yes | No | No |
Oxfendazole | No approvals | ||
Thiabendazole | Yes | Yes | No |
Imidazothiazole | |||
Levamisole | No approvals | ||
Tetrahydropyrimidines | |||
Morantel | Yes | No | No |
Pyrantel | No approvals | ||
Macrocyclic Lactones | |||
Ivermectin | No approvals | ||
Doramectin | No approvals | ||
Moxidectin | No approvals | ||
Eprinomectin | No approvals | ||
Salicylanilide | |||
Closantel | No approvals | ||
Organophosphate | |||
Haloxon | No approvals | ||
This table is based on information from the Food Animal Residue Avoidance Databank (FARAD) obtained May 8, 2023. Label claims regarding specific parasite species or genera as well as susceptible life stages (adult vs. larval vs. inhibited stages) must be observed. Resistance has been reported in all drug classes listed. | |||
The benzimidazole drug class includes albendazole, fenbendazole, oxfendazole, and thiabendazole. These drugs have variable activity against inhibited larvae, such as Ostertagia.
Fenbendazole has a wide margin of safety, and its doses and frequency of administration can be increased above the label amount in many cases (veterinarians must follow laws, requirements, and restrictions for extra-label drug use). Studies on fenbendazole have shown an increased killing capacity with multiple-day dosing because the efficacy of this dewormer increases with increased length of drug exposure.
In contrast, albendazole has a narrow safety margin, and neither the dose nor the frequency can be increased safely. Overdose with albendazole results in teratogenesis, bone marrow suppression with pancytopenia, and death. Fasting animals before administering benzimidazoles slows GI transit time and increases exposure of the parasite to the anthelmintic, potentially increasing its efficacy. Resistance to benzimidazoles is not uncommon, and once resistance has developed, susceptibility is not regained, even when the drug is not administered for several years on a farm.
Levamisole is an imidazothiazole drug and has a narrow therapeutic index because it targets acetylcholine receptors and can affect those of the host as well as the parasite; however, it is water soluble and easy to administer. Levamisole has very little activity against inhibited larvae. Morantel and pyrantel are tetrahydropyrimidines and have a wider margin of safety than levamisole. Morantel is a feed additive and, like other any anthelmintic delivered in feed, may not be administered in an effective dose if the animal does not consume the entire amount. Underdosing may result and, eventually, lead to anthelmintic resistance.
Signs of toxicosis from imidazothiazole or tetrahydropyrimidine drugs include typical “SLUD” clinical signs—salivation, lacrimation, urination, and defecation—in addition to hyperexcitability, trembling, ataxia, collapse, and death from respiratory failure.
The macrocyclic lactone drug class includes the drugs ivermectin, doramectin, moxidectin, and eprinomectin. These drugs are highly effective against adult, larval, and inhibited larval stages.
When first introduced on the commercial market, macrocyclic lactones were able to control both internal and external parasites, were inexpensive, and were highly effective. Because they were used heavily in suppressive deworming programs after being introduced, resistance to the macrocyclic lactones is now widespread in GI nematodes around the world. Pour-on formulations may have extended activity, thus decreasing labor costs associated with application; but because concentrations wane over a longer period, parasites may be exposed to sublethal doses, potentiating the development of anthelmintic resistance.
All drugs are available in a variety of forms (oral, injectable, and pour-on) to suit all types of management systems. Outside of the US, several additional drugs are available in each drug class, including combination deworming products. Some narrow-spectrum anthelmintics, such as the salicylanilide closantel, have excellent activity against Haemonchus contortus in sheep and remain in the host for a long time. Haloxon, an organophosphate, is approved for use against GI nematodes in calves.
Cattle: Special Considerations
GI parasitism is most common in young beef cattle from the time of weaning and several months after, and in segregated groups of dairy calves during the first grazing season. Immunity to GI nematodes is acquired slowly; two grazing seasons may be required before a non-sterilizing protective immunity is attained. In endemic areas, cows may continue to harbor low burdens, which may contribute to suboptimal production on some farms. GI parasitism in juveniles may be controlled by the administration of broad-spectrum anthelmintics in conjunction with pasture management to limit reinfection, such as alternate grazing with other species or integrated rotational grazing in which susceptible calves are followed by immune adults.
In beef herds, anthelmintic treatment at weaning is of value, particularly if the young cattle are to be retained—eg, as replacement heifer stock or as steers to be fed. Cattle finished on grass should receive treatment at weaning and at intervals throughout the next 12 months and, if possible, should be moved to safe pastures to maximize liveweight gain.
When cattle cannot be moved readily to different pastures, strategic treatments may limit the contamination of pastures and rapid reinfection. Alternatively, intraruminal boluses may be used in countries where approved.
In warm temperate regions of the world, such as Australia and New Zealand, the southern US, and the large cattle-raising regions of southern Brazil, Uruguay, and Argentina, young cattle receive two or more treatments from late summer and into fall to prevent large increases in pasture contamination and infection during winter and spring. Two or three strategic treatments administered at short intervals from the time of weaning in such regions could be just as effective as spring treatments in cool temperate regions. However, because infective larvae tend to survive on pasture beyond the time of fall weaning in warm temperate regions, longer intervals between treatments (eg, at weaning, during the winter, and in late spring) may be more effective.
When type II ostertagiasis is a problem, treatment with an anthelmintic effective against inhibited larvae is recommended before the expected time of outbreak. For more information on the life cycles of specific parasites, see Common Gastrointestinal Parasites of Cattle.
The economic justification for administering dewormers in dairy animals is controversial. The benefit justifies their use in first-calf heifers and replacement stock; however, the use of dewormers in dairy cows that are completely confined or in cows with subclinical GI parasites generally derives little to no benefit in milk production. More informed treatment decisions may become feasible with the use of ELISA to monitor for Ostertagia antibodies in bulk-tank milk.
Small Ruminants: Special Considerations
A special strategic treatment is required in most regions to counter the periparturient relaxation of immunity (resulting in the periparturient rise in worm egg output) that occurs in breeding ewes and does. The precise timing of such treatment varies by region and for different species of parasites; in temperate regions, the treatment depends on whether dams and offspring are turned out onto clean or contaminated pasture.
On clean grazing, only the dams (with an existing parasite burden) act as a source of worm eggs and therefore require treatment to prevent pasture contamination and subsequent infection of their offspring. Dams treated during the month before lambing or kidding not only should exhibit a drop in worm egg output but also may show improved productivity. On contaminated pasture, both dams and their offspring pass worm eggs in their feces (dams from their existing worm burden; offspring, from larvae overwintering on pasture).
Sheep and goats are more consistently susceptible to the adverse effects of worms than cattle are, and disease is more common in sheep and goats. Immunity to the parasites is acquired slowly and is generally incomplete. Frequent treatments may be required, particularly during the first year of life, although a good understanding of local parasite epidemiology will ensure that such treatments are appropriately timed. Subclinical goats and sheep have not been shown to benefit (in terms of average daily gain, feed efficiency) from deworming, and this practice should be discouraged. For more information on the relevant life cycles of specific parasites, see Common Gastrointestinal Parasites of Small Ruminants.
Combination Anthelmintic Treatment
Administering full doses of two or three classes of dewormers simultaneously can improve efficacy through an additive effect that resembles the concept of broad-spectrum antibiotic coverage. In addition to management changes, this deworming approach can be useful in herds with documented multidrug resistance and limited deworming options. Because fewer resistant GI nematodes survive and a larger refugium provides greater dilution of the resistant gene pool, combination deworming can even slow the development of multidrug resistance. Meat or milk withdrawal times should be based on the drug that has the longest withdrawal interval.
Anthelmintic-related Diagnostic Techniques
The increase of severe, widespread multidrug resistance has necessitated the monitoring of drug efficacy. Drug resistance should be suspected when other factors, such as improper dosage, poor nutrition, rapid reinfection, or disease other than GI parasites, are excluded. The fecal egg count reduction test (FECRT) is the primary means of assessing drug efficacy within a herd.
The FECRT should be conducted in individual herds every 2–3 years during the time of year when egg counts are expected to be the highest, as suggested by the local epidemiology of parasites in the region. Ideally, 10–15 younger animals with higher anticipated egg counts or FAMACHA scores should be chosen for this test, and an untreated control group can be used to account for egg count fluctuations unrelated to a drug treatment. For very small flocks or herds, results from an FECRT should be interpreted with caution because individuals can skew the egg counts more powerfully in small groups.
Fecal egg counts (FECs) are conducted on the same day that an anthelmintic is administered; then a second fecal egg count is made on the same individual between 10 and 14 days after application of the treatment. The time period between the pretreatment and posttreatment samples is critical to avoid confounding with reinfection by a separate parasite population. The fecal egg count reduction (FECR) can be calculated as follows: %FECR = ([pretreatment FEC − posttreatment FEC]/pretreatment FEC) × 100.
See current recommendations on the use of FECRT from the World Association for the Advancement of Veterinary Parasitology.. FECRT trends over time enable more informed decision-making about drug treatments.
Refugia of GI Parasites in Ruminants
Practices that lead to a larger protective refugium include targeted selective treatment, selective nontreatment, or whole-herd targeted treatment during high-risk periods. Parasites are not evenly distributed in a herd. Most of the parasites (75%–80%) are often in a small portion (20%–25%) of the animals, and these animals contribute the most parasite eggs to pasture contamination. If these animals can be identified with FAMACHA scoring, body condition scoring, production metrics, and/or fecal testing, targeted treatment can eliminate this large source of pasture contamination without necessitating treatment of the entire herd.
In targeted selective treatment, only animals that would benefit from the administration of an anthelmintic are treated, including animals that have disease or decreased production metrics.
In selective nontreatment, a portion of the herd that appears not to require an anthelmintic is purposely left untreated.
Whole-herd targeted treatment can be optimized to particular high-risk periods on the basis of monitoring and diagnostics (epidemiologic factors, grazing history, nematode species present in the herd). For example, data spanning a couple of years might lead a cow-calf operation to optimize its deworming protocol to two treatments per year.
Differences in management systems may make one of these options more feasible than another; however, any of these three options will provide a larger refugium and mitigate the development of drug resistance.
Practices that lead to a smaller refugium and the acceleration of drug resistance include suppressive deworming, rotational deworming, and deworming followed by immediate transfer to a clean (rested) pasture. These practices are no longer acceptable.
Biosecurity and Quarantine of Ruminants with GI Parasites
Animals with anemia, diarrhea, weakness, and depression should be separated from the rest of the herd to facilitate treatment and feeding, avoid bullying by herdmates, and minimize environmental contamination within the herd. Standard biosecurity measures, such as feeding sick animals last and avoiding shared equipment will prevent cross-contamination.
New herd additions or animals that have traveled should be quarantined for 3–4 weeks in a stall or dry lot with no exposure to the native herd. Because most trichostrongyles will begin producing eggs in this time frame, infections can be detected with fecal testing and treated before being introduced to the native herd. Resistant parasites can be unintentionally "purchased" in this manner, so dewormer efficacy should be confirmed with FECRT. If resistance is detected, combination deworming with two or three drug classes may be necessary.
Alternative (Nondrug) Adjunctive Options for Treating GI Parasites in Ruminants
A wide variety of nondrug control measures exist for ruminants and can be used in an adjunctive manner as part of an integrated parasite management plan. Approaches to pasture management include resting pastures to allow for the natural attrition of infectious L3 larvae, managing forage height, and allowing mixed-species grazing.
High amounts of dietary protein enhance the ability of ruminants to resist clinical parasitism and support the immune response to parasite challenge. The administration of copper oxide wire particles as a bolus has been shown to decrease the Haemonchus contortus worm burden in sheep and goats; however, care must be taken to avoid copper toxicity in these sensitive species. The addition of condensed tannin forages and predacious, nematode-trapping fungi (Duddingtonia flagrans) in feed has also been shown to decrease worm burden and fecal egg counts in treated animals; however, these substances need to be fed continuously.
Vaccination of small ruminants and camelids with novel parasite proteins to stimulate a protective immune response has been studied in Australia and Brazil,4, 5 and it may be a viable option in the US if its use becomes approved.
References
Greer AW, Van Wyk JA, Hamie JC, Byaruhanga C, Kenyon F. Refugia-Based Strategies for Parasite Control in Livestock. Vet Clin North Am Food Anim Pract. 2020;36(1):31–43. doi: 10.1016/j.cvfa.2019.11.003. PMID: 32029187
Zajac AM, Garza J. Biology, Epidemiology, and Control of Gastrointestinal Nematodes of Small Ruminants. Vet Clin North Am Food Anim Pract. 2020;36(1):73–87. doi: 10.1016/j.cvfa.2019.12.005. PMID: 32029190
Kaplan RM. Biology, Epidemiology, Diagnosis, and Management of Anthelmintic Resistance in Gastrointestinal Nematodes of Livestock. Vet Clin North Am Food Anim Pract. 2020;36(1):17–30. doi: 10.1016/j.cvfa.2019.12.001. PMID: 32029182
VanHoy G, Carman M, Habing G, et al. Safety and serologic response to a Haemonchus contortus vaccine in alpacas. Vet Parasitol. 2018;252:180–186. doi: 10.1016/j.vetpar.2018.02.014. Epub 2018 Feb 9. PMID: 29559145
Adduci I, Sajovitz F, Hinney B, et al. Haemonchosis in Sheep and Goats, Control Strategies and Development of Vaccines against Haemonchus contortus. Animals (Basel). 2022;12(18):2339. doi: 10.3390/ani12182339. PMID: 36139199; PMCID: PMC9495197
