Specific Diseases of the Cardiovascular System

Heartwater

Heartwater, also known as cowdriosis, is an infectious, non-contagious, tick-borne rickettsial disease of goats, sheep, and cattle, as well as some wild ruminants.

Historically restricted to sub-Saharan Africa, heartwater has now also been identified in the Caribbean.

Etiology

The causative agent is the rickettsia Ehrlichia (previously Cowdria) ruminantium. It is Gram negative and stains red­dish purple to blue in smears with Giemsa stain. Pleomorphism is common. Smaller organisms are usually coccoid, while larger ones may be horseshoe, ring, or rod shaped. In mammalian hosts, the organism has a predilec­tion for vascular endothelial cells. In the tick vector, it is found in intestinal epithelial cells and in cells of the sali­vary glands.

Historically, efforts to perform in vitro cultures of E. rumi­nantium have failed, but improved, chemically defined cul­ture media for propagation of the organism are now available (Zweygarth and Josemans 2001). The organism can also be propagated on various types of endothelial cell lines as well as tick cell lines (Bezuidenhout et al. 1985; Bell-Sakyi et al. 2000). Live organisms in whole blood or tissue homogenates from infected animals may be stored for extended periods by quick freezing in liquid nitrogen using a suitable cryopreservative, for example dimethyl sulfoxide.

There are eight distinct genotypes identified within the species E. ruminantium, based on 16S rRNA gene sequences, and within these various genotypes multiple strains may occur. Cross-protective immunity may develop after infection with different field strains (Van Winkelhoff and Uilenberg 1981; Uilenberg 1983), but strains are often not or only partially cross-protective when used to chal­lenge ruminants (Uilenberg 1983; Jongejan et al.

Based on gene sequencing, an Ehrlichia sp. closely related to E. ruminantium and referred to as Panola moun­tain Ehrlichia has been detected in lone star ticks (Amblyomma americanum) in the state of Georgia in the United States. The infection was found to be transmissible to a goat through feeding by infected ticks, and uninfected nymphal A. americanum ticks that subsequently fed upon the infected goat became infected (Loftis et al. 2006).

Epidemiology

Heartwater is indigenous to sub-Saharan Africa. It was first recognized in South Africa in 1838 and is still considered a major obstacle to the expansion and development of live­stock enterprises on the continent. Imported cattle, sheep, and goats in particular are very susceptible to severe infec­tion, with high mortality rates. Heartwater is also found on islands near Africa, including Madagascar, Mauritius, Reunion, the Comoros, and Sao Tome.

The introduction of heartwater to the Western Hemisphere causes additional concern. The disease was reported on the island of Guadeloupe in 1980. Since that time, it has been confirmed on two other Caribbean islands, Antigua and Marie Galante. The tick vector Amblyomma variegatum, originally introduced into the region with an importation of cattle from Senegal in the nineteenth or possibly even eighteenth century, has also been found on Puerto Rico, Vieques, St. Croix, St. Martin, Anguilla, St. Kitts, Nevis, La Desirade, Martinique, St. Lucia, St. Vincent, and Barbados, but E. ruminantium has not yet been identified on these islands (Camus et al.

The distribution of heartwater reflects the geographic distribution of the tick vectors of the genus Amblyomma that transmit the disease to domestic and wild ruminants, and occurrence of the disease is linked to tick numbers and feeding activity, with increases noted after heavy rains. Amblyomma spp. are three-host ticks that feed on a wide variety of mammals and birds. The most widespread vector for heartwater in Africa and the Caribbean is A. variega- tum, also known as the tropical bont tick. Other natural African vectors include Amblyomma hebraeum in south­ern Africa, Amblyomma pomposum in south-central Africa, and A. gemma and A. lepidum in east and northeastern Africa. Several other African Amblyomma spp. may carry the organism, but mainly feed on wild animals. At least two ticks found in North and South America - A. macula- tum, the Gulf Coast tick, and Amblyomma cajennense - the cayenne tick, are capable of transmitting E. ruminantium to domestic ruminants, the former quite effectively (Uilenberg et al. 1984).

In general, Amblyomma ticks can passage E. ruminan- tium transstadially but not transovarially, though there is one report of transovarial transfer. Consecutive passage occurs from larval to nymphal stage, nymphal to adult stage, and larval to nymphal to adult stage. Ticks therefore can remain infected for periods as long as several years. The organism resides in the intestinal epithelium of ticks and is thought to be transmitted with the saliva (Kocan and Bezuidenhout 1987).

The hosts for E. ruminantium appear to be essentially members of the family Bovidae, including both domestic and wild ruminants in Africa. Cervidae are also suscepti­ble, including the white-tailed deer (Odocoileus virgin- ianus) in North America. Wild ruminants may serve as a reservoir for infection, but a wildlife reservoir is not neces­sary to sustain infection.

An age-related resistance to disease occurs in young domestic ruminants, independent of maternally derived passive immunity. The period of resistance in calves and lambs gradually wanes after approximately the first three weeks of life, and in kids, after the first two weeks. Young animals exposed during this period are unlikely to develop clinical disease and are resistant to subsequent homolo­gous re-infection. Nevertheless, infections have been detected in kids free of ticks under two weeks of age using PCR techniques, suggesting that vertical transmission from doe to kid occurs, either in utero or through colostrum (Faburay et al. 2007).

Goats are the most susceptible natural hosts, based on epi­demiologic observations and experimental challenge stud­ies. Indigenous goats in endemic areas are more resistant than imported ones, but serious incidents of acute heartwa­ter in local goats do occur in Africa (Aklaku 1980; Gueye et al. 1984). Very young, resistant goats may not be exposed to feeding ticks during their resistant period because of local variations in tick populations, preferential feeding by ticks on cattle over goats, or confinement of kids to villages to avoid theft and predation (Ilemobade 1977). A seropreva­lence survey of heartwater in Red Maasai sheep and Small East African goats conducted at three sites in Narok District of Kenya using the MAP1-B enzyme-linked immunoab­sorbent assay (ELISA) found 62-82.5% of sheep and 42.5-52% of goats to be seropositive (Wesonga et al. 2006).

Evidence for differences in breed susceptibility is sug­gested by epidemiologic and experimental observations. In South Africa, Angora goats are quite susceptible to heart­water compared with other indigenous or exotic breeds (Van de Pypekamp and Prozesky 1987). Breed differences in susceptibility have also been reported in Guadeloupe, with native Creole goats showing greater resistance than European breeds of goats, possibly due to genetic resist­ance associated with a recessive sex-linked gene (Matheron et al.

Pathogenesis

E. ruminantium is introduced into the mammalian host by an infected tick during feeding. The early development of infection is not well clarified. It has been suggested that initial replication occurs in regional lymph nodes within macrophages and other reticuloendothelial cells. This is followed by a rickettsemic phase lasting one to four days accompanied by fever. The organism can be demonstrated to occur in the blood plasma as well as in neutrophils (Logan et al. 1987). Subsequently, the organism invades and multiplies in vascular endothelium throughout the body, particularly in the cortex of the brain. The ensuing vasculitis results in fluid and protein loss through capillar­ies with local edema and hemorrhage that, depending on the location and severity, account for the varied clinical and postmortem findings. The incubation period in goats after experimental intravenous inoculation is 7-14 days, or even shorter after a massive infective dose.

Clinical Findings

The incubation period to fever after natural tick-transmitted infection is usually from about two to as much as four weeks. There are four clinical forms of heartwater: pera­cute, acute, subacute, and subclinical. Their development depends on host susceptibility and virulence of the infec­tive strain. Sporadic cases or epizootic outbreaks may occur. Goats most often experience peracute and acute infections.

In peracute disease, affected goats develop a high fever and then suddenly collapse without warning, followed by a period of convulsions or paddling lasting from minutes to several hours. The acute form may last from two to five days.

Signs of subacute disease may be limited to fever, watery eyes, mucous nasal discharge, coughing, dyspnea, and pos­sibly diarrhea and mild nervous signs. Subacute disease is most likely in previously exposed or naturally resistant animals. Subclinical infection, manifested by a transient fever, is uncommonly recognized in goats.

The author (DMS) observed an outbreak of cowdriosis in goats at a cross-breeding operation in Ethiopia that dra­matically demonstrated the difference in susceptibility of imported and indigenous animals. Imported purebred dairy goats on the premises developed peracute disease and died suddenly. The offspring of cross-bred imported and indigenous-breed goats manifested the acute form of the disease, with fever, cardiac, and neurologic signs, while the indigenous goats on the premises showed subacute or no signs of disease.

Clinical Pathology and Necropsy

Decreases in packed cell volume, hemoglobin, total plasma protein, and serum albumin are common findings in heart­water. The leukocyte response is variable in goats, with either neutropenia and lymphopenia or lymphocytic leu­kocytosis reported (Ilemobade and Blotkamp 1978; Abdel Rahim and Shommein 1978). Hyperglycemia and lactic acidosis may be recorded terminally. Orange-yellow serum has been reported as a consistent finding in Angora goats affected with heartwater, but the observation has not been repeated in other breeds of goats.

Gross postmortem findings in goats include hydroperi­cardium, variable degrees of hydrothorax, and pulmonary edema. Other possible signs are ascites, edema of lymph nodes, serosal hemorrhage particularly of the heart, mucosal congestion of the gastrointestinal tract, and swol­len kidneys. Splenomegaly, a common sign in other spe­cies, was not observed in experimentally infected goats (Ilemobade and Blotkamp 1978). Severe nephrosis is reported in Angora goats (Prozesky and Du Plessis 1985).

Histologically, the disease is characterized by the pres­ence of clusters of E. ruminantium in the vascular endothe­lium of virtually all tissues examined; brain cortex is the most reliable source and liver the least reliable.

Diagnosis

Diagnosis is presumptive, based on the clinical history and signs, the presence of Amblyomma ticks in the region and on the animal, and the demonstration of organisms in vas­cular endothelium on smears or histologic examinations, as described above. Differential diagnosis for the peracute form of disease includes virtually all causes of sudden death in goats, discussed in Chapter 16. The acute or neu­rologic form of heartwater must be distinguished from tet­anus, rabies, pseudorabies, organophosphate toxicity, and various plant poisonings. In particular, one leguminous tree, Albizia versicolor, occurs in areas of southern Africa where heartwater is endemic. Consumption of the seed pods by goats can produce both neurologic signs and hydropericardium, which are suggestive of heartwater (Soldan et al. 1996). When diarrhea and fever precede neu­rologic signs, peste des petits ruminants and salmonellosis should be ruled out.

Confirmation of heartwater in live animals was histori­cally difficult because the organism was not easily cultured in vitro. Currently it can be isolated from the blood of an infected host using cultivation on ruminant endothelial cells. Various polymerase chain reaction (PCR) techniques also are now available and are preferred for confirmation of clinical cases. These include real-time PCR, nested PCR, and multipathogen real-time PCR (OIE 2018).

In the field, diagnosis in dead animals can be made through preparation of smears of brain gray matter at nec­ropsy to identify colonies of E. ruminantium in capillary endothelial cells of the brain tissue. Cerebral cortex can be obtained by drilling or punching a hole in the skull, or cere­bellar cortex can be obtained with a spatula through the foramen magnum after removing the head. Proper biosecu­rity should be observed during this procedure in places where rabies is endemic. A piece of gray matter the size of a match head is placed on a microscope slide, crushed to a paste consistency by another slide, and, while maintaining pressure, drawing the slides over each other lengthwise to produce a single layer of cells. The slides should be air­dried and fixed in methanol for subsequent staining with eosin and methylene blue or Giemsa (OIE 2018). Microscopic examination will reveal blue- to purple-stained clusters of E. ruminantium in vascular endothelium.

In general, serologic tests for detecting antibodies to E. ruminantium all suffer to some extent from lack of speci­ficity, as there are cross-reactions with some other Ehrlichia species or unknown agents. This has partly been overcome by using more specific recombinant antigens instead of crude antigens for ELISA (Kakono et al. 2003). Currently there is an indirect ELISA that uses a recombinant antigen expressed as a partial fragment of the major antigenic pro­tein 1 (MAP1), called MAP1-B, which gives improved spec­ificity over earlier methods, as does a competitive ELISA that uses the map1 gene cloned in a baculovirus and mono­clonal antibodies raised against the MAP1 protein, MAP1-C. However, both assays still detect cross-reacting antibodies to other Ehrlichial organisms, including Ehrlichia chaffeensis, E. canis, and Panola mountain Ehrlichia (OIE 2018), and are therefore not especially use­ful in serosurveillance, as has been confirmed in a study conducted in the Caribbean region (Kelly et al. 2011). This lack of specificity would be of particular concern in North America, where E. ruminantium does not yet occur, but Panola mountain Erlichia is already present. Notably, the development has been reported of a dual-plex Taqman™ (Applied Biosystems, Waltham, MA, USA) qPCR assay tar­geting the groEL gene of Panola mountain Erlichia and E. ruminantium that can distinguish infection between the two species (Sayler et al. 2016).

Treatment

Successful treatment depends on early initiation of antibi­otic therapy. Animals treated during the incubation or febrile stage of acute heartwater respond favorably to tetra­cycline, 10-20 mg/kg bw intravenously or intramuscularly, administered at the first sign of fever and repeated one additional time 24 hours later. Long-acting oxytetracycline given once intramuscularly at the onset of fever at a dose of 20 mg/kg bw is also effective. In some cases goats, as well as cattle and sheep that initially respond well to treatment, may show a recurrence of fever 20-25 days post treatment. In severe cases, they should be retreated at that time. Therapy initiated after the onset of neurologic signs is almost always ineffective.

Antibiotics can also be considered for prophylactic use to manage heartwater, when susceptible exotic breeds are imported and introduced to endemic areas. In goats, short­acting oxytetracyclines at a dose of 3 mg/kg bw on days 10, 20, 30, 45, and 60 after their introduction is recommended, with animals not dipped for ticks until day 60 (Gruss 1981).

Control

Attempts to control heartwater by controlling tick popula­tions in Africa have been somewhat successful, though eradication of the disease by vector control seems unlikely. Coordinated attempts to eradicate the A. variegatum vector from the Caribbean islands have not succeeded.

Strategic use of acaracides on animals through dipping can be helpful by reducing tick burdens. The intention is not to eliminate ticks altogether, but to achieve a host-vec­tor equilibrium that allows livestock to maintain immunity to heartwater and other tick-borne diseases. The costs of acaracides and their application as well as the potential for development of acaracide resistance must be considered. It is advised that dipping of Angora goats during pregnancy or cold weather should be avoided, because stress-related abortion and hypothermia are common in this breed (Gruss 1987).

Vaccination has long been considered the best solution for the effective control of heartwater, but the develop - ment of a reliable commercial vaccine remains elusive, despite concerted efforts to develop killed vaccines, atten­uated live vaccines, and recombinant vaccines. A detailed description of these efforts is available (Allsopp et al. 2018). A key challenge is the antigenic variability of different strains of E. ruminantium and the failure of vaccine strains to cross-protect. In lieu of an effective vac­cine, a combination of controlled exposure and treatment has been used to vaccinate exotic livestock in Africa, with some success.

Infective materials used as vaccines are prepared by snap freezing of blood (or tissue preparations) derived from arti­ficially infected livestock (or ticks). Because these are rather crude preparations, anaphylactic reactions can and do occur in a small percentage of vaccinates, and other pathogens may unintentionally be transmitted. For vacci­nation these substances are administered by the intrave­nous route. At the first sign of fever, vaccinated animals are treated with oxytetracycline (10 mg/kg intramuscularly or intravenously) to diminish the signs of disease and impart an immunity to re-infection.

Kids vaccinated during their period of natural resistance may not show a febrile response (Van der Merwe 1987). Because of the logistic difficulty of determining the onset of fever in individual animals and recording temperatures, block treatments on a predetermined date after vaccination have been recommended. The time of onset of the febrile response depends on the strain and preparation of the vac­cine. When Angora goats were monitored after vaccination with a frozen Ball 3 strain vaccine, 97% were in the febrile phase between days 10 and 14, while only 76% of goats given a fresh Ball 3 strain vaccine were febrile during that period (Erasmus 1976). Because successful vaccination depends on proper timing of treatment, the mean incuba­tion period for the vaccine to be used should be known in advance if such block treatment is to be used. It may be several days shorter when tick filtrate vaccines are used. It is recommended that temperatures be taken in the early morning and should reach 39.5 °C (103.1 °F) in goats before treatment.

The duration of immunity in goats is not well known, but may be as short as two months. Field exposure to anti- genically different strains may be responsible for a seem­ingly short duration of immunity, and homologous immunity is likely to be longer. Vaccination should be timed to precede periods of peak tick feeding activity in wet seasons.

Schistosomosis

Schistosomes are trematode parasites of the vascular system. Different species reside in the vasculature of different organs, and can produce a variety of clinical manifestations including rhinitis, enteritis, hepatitis, or pneumonia. All clinical forms occur in goats in endemic regions of the world. Livestock, including goats, can serve as a reservoir of infection for schistosomosis in humans.

Etiology

In the family Schistosomatidae are two genera that can cause clinical disease in goats: the Schistosoma and the Orientobilharzia. Numerous species use goats as a definitive host. These are identified in Table 8.3 along with their geo­graphic distribution, intermediate and definitive hosts, and the site of localization within the goat. There is one report of finding Schistosoma incognitum in goats in a slaughter­house survey in Jabalpur, India (Agrawal and Sahasrabudhe 1982), though this species is usually associ­ated with swine. Experimental patent infection of goats with S. incognitum was later demonstrated (Gupta and Agrawal 2005). Schistosoma curassoni, previously thought to be synonymous with S. bovis, is now considered a distinct species infecting goats, sheep, and cattle (Verycruysse et al. 1984), though hybridization is known to occur between the two species (Rollinson et al. 1990). Ongoing phyloge­netic studies are helping to clarify the relationships and classification of the pathogenic species of Schistosomatidae (Snyder and Loker 2000; Webster et al. 2006).

The schistosomes are elongated trematodes with distinct male and female forms that are found together in the host as mating pairs. The life cycle of schistosomes is indirect and involves aquatic snails as intermediate hosts. Though there are species variations in life cycle, the general pattern is as follows (Soulsby 1982). Adult schistosomes reside in the vasculature of the target organ of their definitive hosts. These organs are usually the liver, intestine, nasal mucosa, or urinary bladder. Gravid females lay eggs that pass through the vessel walls and gain entrance to the gut lumen, bladder lumen, or nasal passage. These eggs, which may already contain live miracidia, hatch when passed into water. The miracidia are released and infect the appropriate species of aquatic snail, which serves as the intermediate host. Subsequent development in the snail occurs over a variable time period of 38-126 days based on environmental factors. Two generations of sporocysts occur in the snail, leading to the formation of cercariae that are released from the snail back into water. When definitive hosts stand in or drink contaminated water, they are infected by the cercar- iae, by penetration of either the skin or the rumen wall.

After entry into the definitive host, cercariae transform into schistosomula and are carried to the lungs and then the liver via the blood stream over a period of one week. Schistosomula are usually present in the portal veins by day eight. Mating occurs in the portal vein and adults then migrate to the mesenteric veins where maturation and egg laying occur. When hyperinfection occurs, adult schisto­somes may mature in the pulmonary vessels and lay eggs in the lung. Pulmonary schistosomosis due to S. indicum has been reported in goats in India (Sharma and Dwivedi 1976). In the case of S. nasale, maturation and egg laying occur in the vessels of the nasal mucosa and eggs are passed in nasal discharge. The prepatent period for S. bovis infection in goats was recorded as 47-48 days (Massoud 1973). The pre­patent period for S. japonicum in goats in China was reported to be 37.7 ± 3 days, and eggs were still present in goat feces 14 months post infection, indicating a prolonged potential for transmission (Shen et al. 2016).

Epidemiology

Schistosomes infect humans and animals mainly in Asia, Africa, the Middle East, South and Central America, and the Mediterranean region. Schistosomosis, or bilharzia, is an important human disease in Africa, Asia, and South America, caused principally by S. haematobium, S. japoni- cum, or Schistosoma mansoni. Livestock, including goats, can serve as a reservoir for Schistosoma spp. infective for humans (Adam and Magzoub 1977).

The intestinal form of schistosomosis in goats is reported most frequently from Asia and Africa. The nasal form, caused by S. nasale, is restricted to the Indian subconti­nent. Economic losses because of schistosomosis in goats result from poor growth and performance, treatment costs, mortality, and condemnations, especially of livers, at slaughter (Singh Nara and Nayak 1972; Seydi and Gueye 1982).

The occurrence of schistosomosis is closely tied to the ecology of the intermediate snail hosts. The snails thrive in stagnant or slowly moving water, such as is found in irriga­tion ditches, marshlands, rice paddies, watering tanks or troughs, shallow ponds or puddles, and ditches during sea­sons of heavy rain. As such, the occurrence of schistosomo- sis may be continuous or seasonal, depending on the nature of the contaminated water source. Where occurrence is seasonal, snails persist through periods of decreased water habitat, mostly through a strategy of prolificacy. Livestock are infected when they drink from, stand in, or wallow in such water sources. Additional factors contributing to an increased prevalence are poor grazing, limited water sup­plies, and overcrowding (Hurter and Potgieter 1967).

The prevalence of schistosomosis in various livestock is largely dictated by their behavior relative to water. Pigs and buffalo, the wallowing species, have a high prevalence of infection; cattle an intermediate prevalence; and sheep and goats a low prevalence (Agrawal 1981). Goats show a dis­tinct aversion to immersion in water, and even avoid walk­ing through it. This may reduce their potential for exposure (Kassuku 1983). In abattoir surveys, prevalence of schisto­somosis in goats is consistently much lower than in buffalo and cattle (Islam 1975; Kassuku et al. 1986). Nevertheless, serious losses of goats can occur, such as reported from China, where summer rains produce marked increases in intermediate host snail populations (Li 1987). When cattle,

Table 8.3 Schistosomes reported to infect goats.

sheep, and goats were experimentally challenged with S. bovis or S. Japonicum cercariae, the intensity and severity of infection were most profound in goats (Massoud 1973; Chiu and Lu 1974).

Schistosomosis is an important zoonotic disease, particu­larly in tropical and subtropical regions of Africa, Asia, and South and Central America. In humans, it assumes both the intestinal and urogenital forms. An estimated 97.2 mil­lion people were treated for the disease worldwide in 2018, the majority in Africa (WHO 2020). People are infected during routine agricultural, domestic, occupational, and recreational activities, which expose them to infested water. Fecal contamination of water by infected livestock is an important factor in the spread of disease to humans. In China, for example, goats have been identified as playing an important role in the spread of S. Japonicum infection to humans, particularly in marshland areas (Wang et al. 2016).

Pathogenesis

The intestinal form of the disease occurs approximately two months after infection, when adults begin to lay eggs in the mesenteric veins and the spined eggs pass through the intestinal mucosa. This results in injury to all layers of the intestinal wall with hemorrhage and edema, and the formation of microabscesses, granulomas, and progressive fibrosis. These changes lead to diarrhea and dysentery and probably malabsorption, as hypoproteinemia is common. The adult parasites cause phlebitis in the mesenteric ves­sels and are sometimes also found in vessels of the urinary bladder and the pulmonary arteries. As many as 1000 pairs of adults have been counted from the mesenteric veins of goats dying with S. mattheei infection (Hurter and Potgieter 1967).

The hepatic form of the disease is considered to reflect a severe cell-mediated immune response to Schistosoma eggs refluxed back into the portal circulation. Soluble egg anti­gens induce a marked eosinophilic, granulomatous reac­tion, leading to extensive damage to the portal vasculature and subsequently severe fibrosis of the portal triads (Soulsby 1982). In humans especially, this results in portal hypertension, with development of varices and possibly congestive heart failure. In ruminants, these cardiovascu­lar effects are not expressed clinically and liver involve­ment is usually detected only at necropsy. It is reported that O. Curkestanicum infection in goats and sheep leads to chronic debility secondary to hepatic cirrhosis and intesti­nal granuloma formation (Soulsby 1982). Anemia occurs in schistosomosis as a result of hemorrhagic lesions in the intestinal wall from migration of eggs and from blood feed­ing by adults in the vessels.

The nasal form of the disease represents an inflamma­tory reaction to the passage of eggs through the nasal mucosa and, to a lesser extent, the presence of adult schis­tosomes in the nasal vessels. The result is nasal congestion, copious nasal discharge, granuloma formation, and dyspnea.

Pulmonary schistosomosis occurs when the host is chal­lenged with large numbers of cercariae. Maturation of excessive numbers of schistosomes in the liver leads to spread of parasitic emboli back to the lungs. Adult schisto­somes lay eggs in the lung vessels and produce multiple, diffuse, nodular granulomata throughout the lung paren­chyma. Emaciation and respiratory distress result (Sharma and Dwivedi 1976).

Clinical Findings

All ages, breeds, and both sexes of goats are affected. Clinical disease occurs in association with the onset of egg excretion. In the intestinal form, diarrhea, anemia, and emaciation are the cardinal signs. The diarrhea is usually watery, but can be mucoid and/or bloody. Anorexia, dehy­dration, and edema are common accompanying findings. The clinical course is usually weeks to months and can result in death, chronic ill-thrift, or sometimes spontane­ous recovery. Clinically, the presentation may be indistin­guishable from gastrointestinal nematodiasis.

In the nasal form of the disease due to S. nasale, there may be weight loss, snoring, sneezing, copious mucoid or foul-smelling purulent nasal discharge, and dyspnea. In pulmonary schistosomosis, emaciation and dyspnea can occur. The hepatic form of the disease is usually not recog­nized clinically; it is overshadowed by the enteric or pul­monary forms.

Clinical Pathology and Necropsy

Anemia and sometimes eosinophilia may be noted in the hemogram. Hypoproteinemia, hypoalbuminemia, and hypergammaglobulinemia may also be present (Pandey et al. 1976). In experimental S. mansoni infection of goats, there were elevations in serum arginase, AST, and bilirubin, and a depletion of liver glycogen (Adam and Magzoub 1977).

In the field, confirmation of infection requires identifi­cation of schistosome eggs in nasal secretions, nasal scrap­ings, feces, rectal scrapings, or possibly urine. Schistosome eggs are generally larger than nematode eggs. They are elongated and spindle shaped and possess a characteristic terminal spine. Direct smears or sedimentation techniques are preferred to flotation methods for finding these trema­tode eggs, but detection may still be challenging. Microscopic examination of squash preps of rectal mucosa obtained with a bowel forceps was 100% effective in diag­nosing schistosomosis in a group of acutely affected sheep and goats (Hurter and Potgieter 1967). For the diagnosis of caprine hepatic schistosomosis in India, egg-hatching techniques with detection of miracidia were more sensi­tive than either the formol-ether sedimentation concen­tration technique or the alkaline digestion technique for egg detection (Vohra and Agrawal 2006). In chronic cases, egg shedding may be notably reduced and diagnosis using coprologic methods will yield false-negative results. Intestinal scrapings taken at necropsy may reveal schisto­some eggs, and adult flukes may be detected by mincing or macerating the mesentery and submerging it in normal saline for six to eight hours at 37 °C (98.6 °F), after which the saline can be examined for the presence of blood flukes (Vohra and Agrawal 2007). All these limitations suggest the need for reliable immunologic and molecular tech­niques of high sensitivity and specificity for accurate diag­nosis of schistosomosis.

However, serologic tests to date are generally unreliable for confirmation of individual cases or for prevalence stud­ies, mainly because of cross-reactions with other trema­todes, notably Fasciola spp., the amphistomes (rumen flukes), and Haemonchus contortus, which are likely to be present under the same environmental and management conditions as schistosomes. Experimental infections of goats with S. japonicum have been confirmed serologically during the prepatent period using ELISA and immunofluo- rescent antibody techniques (Schumann et al. 1984). A dot ELISA test has been evaluated in experimentally and natu­rally infected goats in India (Vohra et al. 2006). Adjustments necessary to improve the specificity of the test resulted in reduced sensitivity, but nevertheless the test was consid­ered to be potentially useful in the field for conducting prevalence studies in locations where infection status is unknown.

A specific nested PCR assay has been developed to detect S. japonicum infection in domestic animals, including goats, using samples of dried blood on filter paper. The tar­get antigen was detected in S. japonicum flukes and from dried blood samples of infected goats but was not detected in Fasciola flukes or Haemonchus contortus worms Zhang et al. 2017).

At necropsy, the carcass is usually emaciated. Adult schistosomes up to 30 mm in length are most frequently found in mesenteric, portal, intestinal submucosal, and subserosal veins. They may also be found in pulmonary veins and veins of the urinary bladder. The liver may have a grayish discoloration and an uneven surface. The lungs may be enlarged, heavy, brown-black, and rubbery, with multiple grayish nodular foci on the pleura and on cut sur­face when pulmonary schistosomosis is present (Sharma and Dwivedi 1976). A catarrhal enteritis is usually present and granulomatous swellings of the mucosa may be noted, as well as areas of petechial or ecchymotic hemorrhage with accumulation of blood in the intestinal lumen. In the nasal form, large granulomas protruding from the nasal mucosa may be noted on cut section of the nasal passages.

Histologically, lesions are associated primarily with eggs rather than adult schistosomes. Eggs in the liver, lung, intestinal wall, and nasal mucosa induce a marked inflam­matory response, with infiltration of eosinophils, lympho­cytes, and macrophages. Granuloma formation around schistosome eggs is common and the immunohistologic basis of these lesions in goats has been studied (Lindberg et al. 1999). The liver lesion is characterized by fibrosis in the region of the portal triads in advanced cases.

Diagnosis

The definitive antemortem diagnosis of schistosomosis depends on identification of eggs in excretions or by biopsy. The intestinal form of the disease must be distinguished from other causes of diarrhea in association with anemia and emaciation, notably gastrointestinal nematodiasis, coccidiosis, and fascioliasis. The nasal form of the disease must be differentiated from other causes of rhinitis, as pre­sented in Chapter 9. At necropsy, the diagnosis is con­firmed by identification of adult schistosomes in the vasculature, or characteristic histologic lesions.

Treatment

In the past, the few drugs available to treat schistosomosis, such as antimony compounds, had narrow margins of safety and frequently produced toxic effects in goats and sheep. Haloxon was also reported to be effective in goats at a dose of 300 mg/kg against the intestinal schistosome S. mattheei, with no noted side effects (Hurter and Potgieter 1967). Currently, praziquantel is being recommended for rumi­nants at an oral dose of 25mg/kg bw, repeated one time 3-5 weeks later. However, a single oral dose of 60 mg/kg was reported to effectively eliminate S. nasale infection from a goat, with no toxic side effects noted (Anandan and Raja 1987). In addition, a single oral administration of tricla- bendazole at a dose of 10 mg/kg bw is reported to be effective for treatment of goats with S. spindale infection in India (Bhoyar et al. 2012).

Control

Efforts at control are directed at reduction of snail interme­diate hosts and exposure of livestock to infective cercariae. When possible, stagnant water sources should be elimi­nated or fenced off, and piped or running water provided instead. Water tanks and troughs should be emptied and cleaned periodically. When ponds or other stagnant water sources must be used, then snail control by application of molluscicides such as copper sulfate or niclosamide has been employed.

To minimize effects on livestock, goats should be treated with praziquantel timed to peaks of likely incidence, such as two months after heavy rains. Currently there are no vaccines against schistosomosis.