Foot rot in pigs is similar clinically to foot rot in other species and is included here for this reason, although the cause of the disease appears different and in most instances the disease is more analogous to foot abscess.
The majority of cases appear to result from secondary infection of lesions that are traumatic in origin. The most common traumatic lesions are erosions of the sole and wall of the claw that occur in pigs reared on rough, abrasive flooring. By themselves these lesions do not usually produce lameness, unless they are extensive, but when pigs are also reared in dirty conditions, infection and subsequent lameness may occur.
Foot lesions are common in pigs of all ages1,2 and bruising of the sole–heel junction, one of the earliest lesions observed can be seen in piglets less than 24 hours old. If the bruising is severe and further trauma is not prevented necrosis will follow quickly in the feet.3,4 These studies have suggested that the cause may be a combination of factors including trauma, contact dermatitis, and subsequent infection. Wet conditions underfoot may cause maceration of the horn and exacerbate the abrasive effect of the flooring. Foot abscess in neonatal pigs is associated with being reared on woven-wire floors.5,6 Dietary deficiency, especially biotin deficiency, may also result in foot lesions that predispose to secondary infection.
Fusiformis necrophorum, Arcanobacterium (Corynebacterium) pyogenes, staphylococci and an unidentified spirochete have been isolated from affected feet. In an outbreak of the disease on a semi-extensive pig farm, Dichelobacter nodosus and other anaerobic bacteria including Prevotella, Peptostreptococcus, Fusobacterium, Porphyromomonas, Bacteroides, and Eubacterium have also been isolated from affected feet.7
The disease has been reported from several countries and is probably universal in occurrence. A study of the prevalence and distribution of foot lesions in finishing pigs in England found that 94% of pigs had at least one foot lesion.8 The prevalence of the different lesions was: toe erosion (33%); sole erosion (62%); heel erosion (13%); heel flaps (14%); white-line lesions (55%); false sand cracks (24%); and wall separation (11%). The hind feet are more commonly affected than the front feet, and on each foot the lateral digits were significantly more frequently affected than the medial digits. Sole erosions, heel flaps, wall separation, and false sand cracks were observed more frequently on the lateral than the medial digit.
Erosive lesions on the foot are common and have been reported at an incidence as high as 65%. They have been reproduced experimentally and the nature of the flooring has a marked influence on claw wear in pigs. Recently poured alkaline concrete and poorly laid concrete with constituents leading to a rough abrasive surface lead to a high incidence. A slope inadequate to allow proper drainage may also be an important predisposing factor. All ages of pigs are susceptible, but clinical lameness is uncommon. In individual herds where the unfavorable predisposing factors prevail, a high incidence of infection and clinical lameness can occur. The disease may cause reproductive inefficiency due to reluctance to stand or mount for mating.
Perforation of the horn leads to infection of the sensitive laminae. The infection may track up the sensitive laminae to the coronary band and discharge to the exterior. Elastolytic activity is a virulence factor involved in the pathogenesis of foot rot in pigs associated with Dichelobacter nodosus and Prevotella melaninogenica.7
Where the disease is due to abrasion of the horn by rough concrete surfaces, a number of characteristic lesions occur, including:
1. Erosion of the sole at either the toe or the heel
2. Bruising of the sole with hemorrhagic streaks in the horn
3. Separation of the hard horny wall from the heel or sole to produce a fissure at the white line, or
4. A false sand crack in the posterior third of the lateral wall of the claw.
In the majority of cases these do not produce lameness nor do they have any apparent effect on productivity. However, when they are extensive or where infection has occurred severe lameness is apparent. In most cases only the lateral digit of one foot is affected. Heat and obvious pain with only moderate pressure being applied to the affected claw are constant findings. Necrosis extends up between the sole and sensitive laminae and may discharge at the coronet, causing the development of a granulomatous lesion, or it may extend to deeper structures of the foot with multiple sinuses discharging to the exterior. A minimal amount of purulent material is present. Productivity is affected with this type of lesion. With deeply infected feet the recovery rate is only fair with treatment. A permanently deformed foot may result and destruction of the pig may be necessary in severe cases. Secondary abscessation in other parts of the body is an occasional sequel and may result in partial carcass condemnation.
Foot abscesses in neonatal pigs are characterized by necrotic pododermatitis, severe osteomyelitis, arthritis, and tenosynovitis.6 The primary sites of injury are located at either5 the point of the toe at the white line,6 the bulb of the heel7, or the haired skin around the coronet, including the interdigital area. The least severe lesions are superficial abrasions or ulcerations of the hoof wall, heel bulb, or interphalangeal skin, with only minimal inflammatory changes in deeper tissues. The most severely affected digits have focal superficial abscesses, or deep, diffuse, purulent inflammation and fibrosis around tendons, joints, and bones.6 The hind limbs are more commonly affected than the forelimbs, and in the hind limbs the medial claws are most likely to have lesions, whereas in the forelimbs the lateral claws are more likely to be affected. Approximately 6% of piglets develop foot abscess prior to weaning. About one-third of litters may be affected and most litters have only one or two affected pigs. Discharge of pus from the coronary band is common and the horny claw may slough, leaving sensitive laminae of one or more claws or accessory digits exposed. Skin necrosis may be present over the carpi, the fetlocks, hocks, coronary bands, and elbows in about 75% of pigs during the first week of life.
Bacteriological examination of discharges from the lesions may aid in deciding the treatment to be used. In foot abscesses of neonatal pigs, bacteria isolated include:
Necrosis of the laminar tissue with indications of progression from an infected sole are the usual findings.
Most other causes of lameness in pigs are not manifested by foot lesions. In adult pigs housed indoors, an overgrowth of the hoof may occur and be followed by underrunning of the sole, necrosis and the protrusion of granulation tissue causing severe lameness and often persistent recumbency. The general appearance of these feet is not unlike that of canker in horses. Swelling of the hoof is caused by an extensive fibrous tissue reaction. Vesicular exanthema and foot-and-mouth disease are characterized by the presence of vesicular lesions on the coronets and snout.
There are few published reports of treatment of foot rot in pigs. Broad-spectrum antimicrobials or penicillin given parenterally seems rational and the use of Nuflor was said to be a successful treatment.9
Prevention of excessive wear of the feet by the use of adequate bedding and less abrasive flooring in pig pens is suggested as a reasonable control measure. Any existing dietary deficiency should be corrected. Of particular interest is the response to biotin supplementation of the diet of pigs in the prevention of foot lesions of various kinds.
1 Penny RHC, et al. Vet Rec. 1963;75:1225.
2 Penny RHC, et al. Vet Rec. 1965;77:1101.
3 Penny RHC Fellowship Thesis, Royal College of Veterinary Surgeons, London, 1968.
4 Penny RHC, et al. Aust Vet J. 1971;47:529.
5 Gardner IA, Hird DW. J Am Vet Med Assoc. 1994;204:1062.
6 Gardner IA, et al. J Am Vet Med Assoc. 1990;196:1791.
7 Piriz S, et al. Vet Rec. 1996;139:17.
Oral infection principally in calves less than 3 months old. Laryngeal involvement in older animals up to 18 months of age
The term ‘oral necrobacillosis’ is applied to infections of the mouth and larynx with Fusobacterium necrophorum. It includes calf diphtheria, in which the lesions are largely confined to the larynx and pharynx, and necrotic stomatitis, in which the lesions are restricted to the oral cavity. They are considered together because the essential lesion and infection are the same in both instances.
F. necrophorum is present in large numbers in the lesions and is considered to be the causative agent, probably aided by prior injury to the mucosa. In the case of the laryngeal disease, the point of entry is thought to be contact ulcers in the mucosa caused by repeated closure of the larynx.1 Both F. necrophorum subsp. necrophorum (biovar/biotype A) and F. necrophorum subsp. funduliforme (biovar/biotype B) are associated with the disease.
The disease has no geographical limitations but is more common in countries where animals are housed in winter or maintained in feedlots. In the United States, infections involving the pharynx and larynx appear to be more prevalent in the western states than in other sections of the country. It is a common disease in feedlots in yearling cattle, often in company with papillomatosis of the larynx.1
There is also a difference in age incidence, necrotic stomatitis occurring mainly in calves 2 weeks to 3 months of age, while laryngeal infections commonly affect older calves and yearlings. Although the disease is more common in housed or penned animals, it can occur in animals running at pasture.2-4
The disease is seen commonly only in cattle but has been observed in sheep and goats.5,6 Laryngeal chondritis has been described in Texel sheep, which may be predisposed to the disease because of anatomical factors, namely the short head of the breed.7 This may affect the shape of the larynx or its relationship to adjacent tissues.
The causative bacterium is a common inhabitant of the environment of cattle and under unsanitary conditions the infection may be spread on dirty milk pails and feeding troughs. Entry through the mucosa is probably effected through abrasions caused by rough feed and erupting teeth. The difficulty of reproducing the disease and the irregularity of its occurrence, even when F. necrophorum is known to be present, suggests the possibility of etiological factors presently unknown.
F. necrophorum is a normal inhabitant of the oral cavity and causes inflammation and necrosis following injury of the mucosa of the oral cavity, pharynx, and larynx. Edema and inflammation of the mucosa of the larynx results in varying degrees of closure of the rima glottidis and inspiratory dyspnea and stridor. The presence of the lesion causes discomfort, painful swallowing and toxemia. Extension of the lesion to the arytenoid cartilages will result in laryngeal chondritis.7 Involvement of the cartilage will usually result in delayed healing or failure to recover completely.
In describing the clinical findings, a distinction must be made between calf diphtheria characterized by involvement of the larynx and the more common necrotic stomatitis. In the former, a moist painful cough accompanied by severe inspiratory dyspnea, salivation, painful swallowing movements, complete anorexia, and severe depression are the characteristic signs. The temperature is high (41°C; 106°F), the pharyngeal region may be swollen and painful on external palpation, and there is salivation and nasal discharge. The breath has a most foul rancid smell.
Examination of the pharynx and larynx by visual inspection through the oral cavity with the aid of a speculum positioned over the base of the tongue will often reveal the lesions. Visual inspection of the larynx is relatively easy and simple with the aid of a cylindrical plastic speculum placed over the base of the tongue in calves and adult cattle. The larynx can be viewed directly and illuminated with a strong source of light. A flexible fiberoptiscope is also useful when available and is necessary for examination of the equine larynx. The mucosa of the larynx and glottis are usually edematous, inflamed and a necrotic lesion is usually present and visible on one or both arytenoid cartilages. The opening of the larynx is commonly reduced due to the edema and inflammation. Careful visual inspection of the larynx during inspiration may reveal that the lesion extends into one or both vocal cords. The examination usually causes considerable discomfort, anxiety and the production of purulent or blood-stained saliva.
Death is likely to occur from toxemia or obstruction to the respiratory passages on days 2–7. Most affected calves die without treatment but only a small proportion of calves in a group are usually affected. Spread to the lungs may cause a severe, suppurative bronchopneumonia.
In calves affected with necrotic stomatitis, there is usually a moderate increase in temperature (39.5–40°C; 103–104°F), depression, and anorexia. The breath is foul and saliva, often mixed with straw, hangs from the mouth. A characteristic swelling of the cheeks may be observed posterior to the lip commissures. On opening the mouth this is found to be due to a deep ulcer in the mucosa of the cheek. The ulcer is usually filled with a mixture of necrotic material and food particles. An ulcer may also be present on the adjacent side of the tongue and cause severe swelling and protrusion of the tongue. In severe cases the lesions may spread to the tissues of the face and throat and into the orbital cavity. Similar lesions may be present on the vulva and around the coronets, and a spread to the lungs may cause fatal pneumonia. In other cases death appears to be due to toxemia.
Bacteriological examination of swabs from lesions may assist in confirming the diagnosis.
Severe swelling, due to edema and inflammation of the tissues surrounding the ulcer, is accompanied by the presence of large masses of caseous material. Occasionally, lesions similar to those in the mouth, pharynx, and larynx may be found in the lungs and in the abomasum. Microscopically, areas of coagulation necrosis are bordered by large numbers of neutrophils and filamentous bacteria.
Necrotic laryngitis is characterized by inspiratory dyspnea and stridor, toxemia, fever, edema, (inflammation) and necrotic lesions of the laryngeal mucosa.
• Neoplasms of the larynx – occur only rarely, usually in mature cattle, and cause chronic inspiratory dyspnea
• Traumatic pharyngitis– may resemble laryngitis but the lesions are obvious on visual inspection of the pharynx. In chronic cases of traumatic pharyngitis there may be peri-esophageal cavities containing rumen contents
• Foreign bodies – i.e. pieces of wire and small wooden sticks may become lodged in the mucosa of the arytenoid cartilages and cause clinical signs similar to necrotic laryngitis.
The lesions of necrotic stomatitis will usually heal in a few days following debridement of the ulcers, application of a solution of tincture of iodine, and oral administration of sulfamethazine at a dose of 150 mg/kg BW daily for 3–5 d where this is labeled for use in food animals, or parenteral penicillin or broad-spectrum antimicrobials. Therapy should be at least for 5 days and therapy for up to 3 weeks may be necessary.
Successful treatment of necrotic laryngitis is dependent on early recognition and prompt therapy with antimicrobials daily for several days. Corticosteroids may be a beneficial adjunctive therapy, especially to reduce the edema. Tracheostomy may be necessary in some cases to relieve dyspnea. Failure to respond is usually associated with chronic suppurative chondritis, which requires subtotal arytenoidectomy.
Necrotic rhinitis is often confused with atrophic rhinitis. It occurs in young growing pigs and may occur in herds where atrophic rhinitis is present and even in the same pig, but there appears to be no relationship between the two diseases. There are a variety of other conditions of the face of the young pig that can be confused.1 The common occurrence of Fusobacterium necrophorum in the lesions suggests that any injury to the face or nasal or oral cavities may lead to bacterial invasion, especially if the environment is dirty and heavily contaminated.2 The incidence of the disease has diminished in recent years, due probably to a general improvement in hygiene in piggeries but possibly also to the declining occurrence of progressive atrophic rhinitis following vaccination and eradication of P. multocida toxigenic type D.
The lesions develop as a necrotic cellulitis of the soft tissues of the nose and face but may spread to involve bone and produce osteomyelitis. Local swelling is obvious and extensive lesions may interfere with respiration and mastication. Depression of food intake and toxemia result in poor growth and some deaths. Treatment by the local application of antibacterial drugs and the oral administration of sulfonamides is satisfactory in early cases. Oral dosing with sulfadimidine has been effective in young pigs.3 Improvement of sanitation, elimination of injuries and disinfection of pens usually result in a reduction of incidence.
The disease differs from atrophic rhinitis by the presence of oral and facial lesions. Necrotic ulcer in pigs may involve the mouth and face but the lesions are erosive rather than necrotic.
Pseudomonas aeruginosa is an occasional cause of infection in large animals. In cattle, there are occasional cases of generalized infection with Pseudomonas aeruginosa, usually following an attack of mastitis associated with this organism. Systemic invasion in cattle is manifested by fibrinous pericarditis and pleurisy and chronic pyelonephritis. Pseudomonas spp. are an occasional cause of septicemia in foals, septic arthritis, vegetative endocarditis in horses1 and placentitis in mares,2 and is isolated from cases of pneumonia in all large animal species. Infection in the urogenital tract is accompanied by infertility.
Outbreaks of otitis media in suckling calves and in sheep following dipping have been associated with pseudomonas infections.3,4 The association of Pseudomonas spp. with fleece rot in sheep is covered elsewhere.
Pseudomonas spp. are commonly isolated from bacterial corneal ulcers and keratitis that develop after trauma to the corneal epithelium in horses5,6 and the organism promotes rapid liquifaction of corneal stromal proteoglycans. Early cases respond to aggressive topical therapy with tobramycin or gentamicin5 although there has been a significant increase in the resistance of isolates from ulcerative keratitis in recent years.7
Infections with Pseudomonas spp. are notoriously difficult to treat. Clinical isolates from animals are usually sensitive to tobramycin, polymyxin B, carbenicillin, and gentamicin.8
1 Travers CW, et al. J South Afr Assoc. 1995;66:172.
2 Hong CB, et al. J Vet Diag Invest. 1993;5:56.
3 Henderson JP, McCullough WP. Vet Rec. 1993;132:24.
4 Davies IH, Done SH. Vet Rec. 1993;132:460.
5 Sweeney CR, Irby NL. J Am Vet Med Assoc. 1996;209:954.
6 Moore CP, et al. J Am Vet Med Assoc. 1995;207:928.
Dermatitis associated with growth of chromogenic Pseudomonas aeruginosa following prolonged wetting of the skin of sheep
Fleece rot develops as an exudative dermatitis following wetting of the fleece by rain. The growth of toxigenic strains of Pseudomonas aeruginosa is believed to be the major cause of the dermatitis, and the fleece coloration that usually accompanies it, but other Pseudomonas spp. including Ps. maltophilia, have been incriminated in the genesis of the condition.1,2 The enzyme phospholipase C in Ps. aeruginosa is a virulence determinant for this disease.3
The disease is common in most parts of Australia, occurs in South Africa and also in areas of New Zealand. Its occurrence is associated with wet years and in these circumstances the incidence in affected flocks varies from 40–100%.2,4
Fleece rot occurs in sheep only in wet seasons and when the fleece is predisposed to wetting by its physical characters.
Prolonged rainfall, sufficient to wet sheep to the skin for a week, is required for this condition to occur. Young sheep are more susceptible than old, and heritable differences in fleece characters affect the susceptibility of individual sheep. These characters are probably related to the ease with which the skin can be wetted.
The degree of ‘grip’ and body skin wrinkling are unimportant as factors affecting susceptibility but fleece weight, fiber diameter and variability, staple density and neck wrinkling are positively correlated with susceptibility.5,6 These characteristics produce visible differences between fleeces. Resistant sheep have closely packed elliptical wool staples with blocky tips and even crimp. Susceptible fleeces have thin staples of unevenly crimped wool and with a fringe-tipped appearance due to the protrusion of thicker wool fibers above the top of the staple. This fringed appearance is visible along the back and sides. Susceptible flocks are characterized by fleeces with longer, heavier, thicker staples with lower crimp frequency and higher fiber diameter and variability.6
Fleeces with a high wax content are less susceptible probably because of the waterproofing effect of the wax. This view is supported by the observation that disruption of the sebaceous layer on the skin increases its susceptibility to wetting.7
Greasy fleece color has been found a good predictor of susceptibility to fleece rot in some studies4 but not others.5 Wool with a high suint content is highly susceptible.8
The disease can be reproduced experimentally by inoculating Ps. aeruginosa epicutaneously and wetting the fleece.9
With prolonged wetting, the conditions of high humidity in the fleece microenvironment, and the availability of rich nutrients from serous exudates and indigenous suint, allow the proliferation of opportunistic skin and fleece bacteria including Ps. aeruginosa, and result in dermatitis. The predominant bacterium is usually Ps. aeruginosa,10 which inhibits the growth of other bacteria and its pyocyanin produces a green color. Its rapid growth is accompanied by the production of the dermonecrotic toxin phospholipase C, which exacerbates the dermatitis and initiates the inflammatory cascade that draws neutrophils and lymphocytes into the skin.11
In the experimental disease there is outpouring of serous exudate and infiltration of leukocytes into dermis but Ps. aeruginosa is localized as aggregates at the leading front of the seropurulent exudate and never penetrates the dermis.12
Other discolorations may occur depending upon the predominance of a particular chromogenic bacterium; many which belong to Pseudomonas spp. Ps. maltophilia can result in yellow coloration, and Ps. indigofera, blue coloration.
The odor produced by the bacteria and the serum protein on the skin surface is very attractive to blowflies, and most body strikes are due to pre-existing fleece rot lesions. To add a further complication, Ps. aeruginosa also proliferates in the presence of organophosphorus insecticides and facilitates its biodegradation.13
Lesions occur most commonly over the withers and along the back. In active cases, the wool over the affected part is always saturated and the tip is more open than over unaffected areas. The wool is leached and dingy and in severe cases can be plucked easily. The skin is inflamed and serous exudate produces bands of matted and colored fibers across the staple. The coloration of the fibers is commonly green, but may be yellow, yellow-brown, or red-brown1 and occurs in fibers at skin level or extending the full length of the staple.
The general health of the sheep is unaffected in typical fleece rot but severe ulcerative dermatitis with mortality associated with Ps. aeruginosa can occur.
A chronic ulcerative and necrotic dermatitis associated with Ps. aeruginosa, occurring on the tail, udder, and legs of sheep and accompanied by green coloration of the surrounding fleece is recorded following excessive rain in the Mediterranean climate zone of Israel.14
Autopsy examinations are not carried out and laboratory examination of the living animal is not usually necessary.
There are differences in the inflammatory response and in peripheral blood lymphocyte subsets between fleece rot-resistant and susceptible sheep.15,16
Fleece rot resembles mycotic dermatitis in body distribution and predisposing factors but the typical scab is not present in fleece rot.
Treatment is unlikely to be of value but some degree of control may be effected by selection of fleece rot-resistant sheep for use in susceptible localities. In these same localities, shearing before the wet season should facilitate drying of the fleece and lessen susceptibility. The heritability of resistance to fleece rot has been estimated to be between 0.35 and 0.4 and selective breeding programs have been advocated11 and genetically selected lines show increased resistance in high-risk environments.17
Chemical means of drying the living fleece have been shown to reduce wetness, fleece rot and blowfly strike. A mixture of zinc and aluminum oxides with sterols and fatty acids (the mixture identified as B26), applied at the rate of 100–200 mL per sheep as a mist-like simulated rain, caused significant reduction in fleece moisture for 10–12 weeks and this could be extended by further applications. Fleece rot was reduced by 60% and blowfly strike by 75%.
A vaccine containing killed Ps. aeruginosa has protected against the severe exudative form of fleece rot1 and there is hope that a vaccine prepared against highly conserved outer membrane antigens of the organism may give more universal protection.12
1 Burrell DH. Adv Vet Dermatol. 1990;1:347.
2 Kingsford NM, Raadsma HW. Vet Microbiol. 1997;54:275.
3 Chin JC, Watts JE. J Gen Microbiol. 1988;134:2567.
4 Keown AM, Reid TC. Proc NZ Soc Anim Prod. 1997;57:83.
5 Cottle DJ. Aust J Agric Res. 1996;47:1213.
6 Raadsma HW. Aust J Agric Res. 1993;44:915.
7 Pascoe L. Aust J Agric Res. 1982;33:141.
8 Lipson M, et al. Aust J Exp Agric. 1982;22:168.
9 Gogolewski RP. Aust J Agric Res. 1996;47:917.
10 Chin JC, Watts JE. Vet Microbiol. 1992;32:63.
11 Chin JC, Gogolewski RP. Wool Technol Sheep Breed. 1991;39:112.
12 Chin JC, et al. Vet Microbiol. 1995;43:21.
13 Merritt GC, et al. Aust Vet J. 1981;57:531.
14 Yerahum I, et al. J Vet Med A. 1995;42:35.
15 Chin JC, Gogolewski RP. Wool Technol Sheep Breed. 1992;39:112.
Ubiquitous soil saprophyte endemic to Southeast Asia, northern Australia and the South Pacific. Occurs primarily 20° north and south of the equator. Transmission is by inhalation of contaminated dust and cutaneous abrasion. Primarily a disease of sheep and goats and humans, occasional disease in horses and subclinical infection in pigs
Burkholderia pseudomallei (Pseudomonas pseudomallei, Malleomyces pseudomallei) is the sole cause. There is considerable genetic variability and strains vary in pathogenicity.1
The disease occurs almost exclusively in tropical countries 20° north and south of the equator and is endemic in South East Asia, Asia, and northern and sub-northern areas of Australia. Disease occurs in rodents, rabbits, pigeons, humans, animals in zoological gardens, dogs, cats, horses, pigs, sheep, goats, alpacas, and camels but rarely in cattle.2,3 In domestic animals the disease has occurred in outbreak form in pigs, goats, and sheep in Australia,3,4 in the Caribbean area and in Cambodia, in horses in Malaysia and Iran, in pigs and cattle in Papua New Guinea and Australia, in horses in France in 1976–19785 and in cattle in Argentina.6
In endemic areas the organism is a ubiquitous soil saprophyte and is present in moist soil and waterholes which are the primary reservoirs from which most infections are acquired. A variety of free-living amoebae including Acanthamoeba and Hartmannella spp. are potential hosts to B. pseudomallei.7,8 The majority of cases in livestock are associated with the ‘wet season’ and exposure to surface water and mud. Infection occurs through inhalation, ingestion, in association with skin wounds via contaminated dust particles or water or by insect bites. Infected animals pass the organism in their feces and the disease in rodents runs a protracted course, making these animals important reservoirs of infection.
B. pseudomallei can survive in water at room temperature for up to 8 weeks, in muddy water for up to 7 months, and in soil in the laboratory for up to 30 months.9 The organism can survive in contaminated injectable drugs and has ability to survive for some time in cetrimide 3% and chlorhexidine 0.3% solution.3 Varying degrees of virulence are observed in different strains of the organism but starvation or other conditions of stress appear to increase the susceptibility of experimental animals to infection.
The disease can be produced experimentally in goats, sheep, rats, mice, hamsters, and pigs.10
Humans are at risk for infection within endemic areas and while this can be zoonotic it can also occur without direct animal contact through inhalation. The disease of humans presents with various clinical pictures ranging from asymptomatic state, localized infection such as pneumonia, to acute fatal septicemia.11
Veterinarians and animal owners are at risk from localized or generalized infection from infected animals. Pregnant women handling goats aborting with this infection have risk for infection and abortion.3 Infected areas are often rural in nature and pasteurization of commercially sold milk should be ensured as should condemnation of infected carcasses at abattoirs.
There is initial bacteremia or septicemia and subsequent localization in various organs. Experimentally induced melioidosis in goats is characterized by septicemia with undulating fever, wasting, anorexia, hindlimb paresis, mastitis, and abortion. Necropsy lesions include widely scattered microabscesses after intraperitoneal injection, and a chronic disease with abscesses in the lungs and spleen when the infection is administered subcutaneously.12 In pigs experimental infection results in a generalized chronic infection.10
Signs consist mainly of weakness, respiratory disease and recumbency with death occurring in 1–7 d. In experimentally infected sheep, a severe febrile reaction occurs and is accompanied by anorexia, lameness and a thick, yellow exudate from the nose and eyes. Some animals show evidence of central nervous system involvement including abnormal gait, deviation of the head and walking in circles, nystagmus, blindness, hyperesthesia, and mild tetanic convulsions. The disease is usually fatal. Skin involvement is not recorded.
The syndrome may resemble the acute form as seen in sheep, but more commonly runs a chronic course with abscessation.13 Mastitis is common in infected goats; one study finding mammary infection in 35% of infected goats.3
Disease is usually chronic and manifested by cervical lymphadenitis, but in some outbreaks there are signs similar to those in other species. In such outbreaks slight posterior paresis, mild fever, coughing, nasal and ocular discharge, anorexia, abortion, and some deaths may occur.
The syndrome is one of an acute metastatic pneumonia with high fever and a short course. Cough and nasal discharge are minimal and there is a lack of response to treatment with most drugs. Other signs in horses include colic, diarrhea, and lymphangitis of the legs. Subacute cases become debilitated, emaciated, and develop edema.5 Affected horses may survive for several months. A case of acute meningoencephalitis is described in a horse. The onset was sudden and manifest with violent convulsions.14
The organism is easily cultured and may be isolated from nasal discharges. Injection into guinea pigs and rabbits produces the typical disease. An allergic skin test using melioidin as an antigen,5 a complement fixation test (CFT), and an indirect hemagglutination (IHA) test are available. The IHA test is recommended for screening and the CFT for confirmation in cases of active melioidosis in goats15 and pigs.16 Affected horses may give a positive reaction to the mallein test.17
Multiple abscesses in most organs, particularly in the lungs, spleen, and liver, but also in the subcutis and the associated lymph nodes, are characteristic of the disease in all species. In sheep respiratory infection is common and these abscesses in the lung contain thick or caseous, green-tinged pus similar to that found in Corynebacterium pseudotuberculosis lesions. Lesions in the nasal mucosa proceed to rupture with the development of ragged ulcers. An acute polyarthritis, with distension of the joint capsules by fluid containing large masses of greenish pus and acute meningoencephalitis have been observed in experimental cases.
A high incidence of lesions in the aorta of goats is reported in Australia. Nine out of 43 (21%) goats had aortic lesions at autopsy. Seven of these goats died as a result of a ruptured aortic aneurysm.3
Treatment is unlikely to be undertaken in farm animals because of the nature of the disease and the risk of exposure to humans. Little information is available on satisfactory treatments of melioidosis in farm animals but recommendations for man are available.18 Penicillin, streptomycin, chlortetracycline, and polymyxin are ineffective but in vitro tests suggest that oxytetracycline, novobiocin, chloramphenicol, and sulfadiazine are most likely to be valuable, with oxytetracycline the preferred drug. In horses chloromycetin has been shown to be an effective treatment.5
Prevention involves removing animals from the contaminating source. Water supplies can be chlorinated. This and the elimination of infected animals and the disinfection of premises should be the basis of control procedures. Housed animals can be removed from soil by raising them from the ground on wooden slats, concrete or paved floors.
1 Fushan A, et al. Res Microbiol. 2005;156:278.
2 Bergin JJ, Torenbeeck LR. Aust Vet J. 1991;68:309.
3 Choy JL, et al. Acta Tropica. 2000;74:153-158.
4 Thomas AD. Aust Vet J. 1981;57:146.
5 Loganathan P, Tan S. Kajian Vet Malaysia. 1983;15:74.
6 Conigliaro S, et al. Vet Argentina. 1999;16:285.
7 Inglis TJJ, et al. Epidemiol Infect. 2004;132:813.
8 Chua KL, et al. Infect Immun. 2003;71:1622.
9 Thomas AD, et al. Aust Vet J. 1981;57:535.
10 Najdenski H, et al. J Vet Med B. 2004;51:225.
11 Leelarasamee A. Acta Tropica. 2000;74:129.
12 Thomas AD, et al. Aust Vet J. 1988;65:43.
13 Van der Lugt JJ, Henton MM. J S Afr Vet Assoc. 1995;66:71.
14 Ladds PW, et al. Aust Vet J. 1981;57:36.
15 Thomas AD, et al. Aust Vet J. 1988;65:261.
16 Thomas AD, et al. J Clin Microbiol. 1990;28:1874.
Acute or chronic form, and characterized by pneumonia and nodules or ulcers in the respiratory tract and on the skin. The disease is highly fatal
Burkholderia (Pseudomonas) mallei is the causative organism. It has close genetic and antigenic relatedness to Burkholderia pseudomallei.1,2 Isolates of B. mallei, recovered from three continents over a period of 30 years have identical allelic profiles.1
Glanders is restricted geographically to Eastern Europe, Asia Minor, Asia, and North Africa. It was more widespread but has been eradicated from most countries.3 Glanders was an important disease when there were large concentrations of horses in cities and armies, but now has sporadic occurrence, even in infected areas.
Horses, mules, and donkeys are the species usually affected. Humans are susceptible and the infection is usually fatal. Carnivores, including lions may be infected by eating infected meat and infections have been observed in sheep and goats.
B. mallei is an obligate parasite and is readily destroyed by light, heat, and the usual disinfectants and is unlikely to survive in a contaminated environment for more than 6 weeks.
Infected animals or carriers that have made an apparent recovery from the disease are the important sources of infection. Chronic nodular lung lesions, which have ruptured into the bronchi, infect upper airway passages and nasal or oral secretions. Spread to other animals occurs mostly by ingestion, the infection spreading on fodder and utensils, particularly communal watering troughs, contaminated by nasal discharge or sputum. Rarely the cutaneous form appears to arise through contamination of skin abrasions by direct contact or from harness or grooming tools. Spread by inhalation can also occur but this mode of infection is probably rare under natural conditions.
An experimental model for disease has been reproduced by intratracheal inoculation of horses with cultures of B. mallei.4 Horses showed fever within 24 to 48 hours of challenge followed by the progressive development of signs of respiratory distress with epistaxis and purulent nasal and ocular discharge. On post-mortem there was lymphadenopathy, ulcerative lesions in the nasal septa and pneumonia.
Horses tend to develop the chronic form, mules and donkeys the acute form, but all types of equid and all ages are susceptible. The disease is more likely when animals are in a stressed state from heavy work and animals that are poorly fed and kept in a poor environment are more susceptible.
The stress associated with movement of a large number of horses can precipitate an outbreak with high mortality rates. In the few animals that recover, there is a long convalescence with the frequent development of the ‘carrier’ state. Animals rarely make a complete recovery.
The disease has little current economic importance, although the threat of horse movement reintroducing glanders into countries that have eradicated it is a concern.5
While humans are not highly susceptible, the infection may gain access through skin abrasions to produce granulomatous disease and pyemia. Infection can also occur from inhalation of infectious material. The case fatality is high. Horse handlers in general are at risk and veterinarians conducting postmortem examinations without proper precautions are at particular risk. The organism is identified as a possible agent of bioterrorism.
Invasion occurs mostly through the intestinal wall and a septicemia (acute form) or bacteremia (chronic form) is set up. Localization always occurs in the lungs but the skin and nasal mucosa are also common sites. Other viscera may become the site of the typical nodules. Terminal signs are in the main those of bronchopneumonia, and deaths in typical cases are caused by anoxic anoxia.
There is a high fever, cough, and nasal discharge with rapidly spreading ulcers appearing on the nasal mucosa, and nodules on the skin of the lower limbs or abdomen. Death due to septicemia occurs in a few days.
Three major manifestations are described:
The pulmonary form manifests as a chronic pneumonia with cough, frequent epistaxis, and labored respiration.
In the nasal form, lesions appear on the lower parts of the turbinates and the cartilaginous nasal septum. They commence as nodules (1 cm in diameter), which ulcerate and may become confluent. In the early stages there is a serous nasal discharge which may be unilateral and which later becomes purulent and blood stained. Enlargement of the submaxillary lymph nodes is a common accompaniment. On healing, the ulcers are replaced by a characteristic stellate scar.
The skin form is characterized by the appearance of subcutaneous nodules (1–2 cm in diameter), which soon ulcerate and discharge pus of the color and consistency of dark honey. In some cases the lesions are more deeply situated and discharge through fistulous tracts. Thickened fibrous lymph vessels radiate from the lesions and connect one to the other. Lymph nodes draining the area become involved and may discharge to the exterior. The predilection site for cutaneous lesions is the medial aspect of the hock, but they can occur on any part of the body.
Animals affected with the chronic form are usually ill for several months, frequently showing improvement but eventually either dying or making an apparent recovery to persist as occult cases.
Disease is accompanied by a low hemoglobin content of the blood, a low erythrocyte count and packed cell volume, and a moderate leukocytosis and neutrophilia. The principal tests used in the diagnosis of glanders are the mallein test, the complement fixation test on serum, and demonstration of the organism.
The intradermopalpebral test has largely displaced the ophthalmic and SC tests. Mallein (0.1 mL) is injected intradermally into the lower eyelid with a tuberculin syringe. The test is read at 48 h, a positive reaction comprising marked edema of the lid with blepharospasm and a severe, purulent conjunctivitis.5 Some infected animals exhibit a general hypersensitivity reaction after inoculation.
The CFT is the most accurate of the serological tests available, and the usual official test, but some strains of B. mallei give cross-reactions with B. pseudomallei. Other tests used are an indirect hemagglutination test using mallein as the antigen6 and the conglutinin complement absorption test.7 The accuracy of the complement fixation test may be improved by the simultaneous testing with the indirect hemagglutination test. CFT may be unsuitable in mules and asses because of anticomplement activity. A dot enzyme-linked immunosorbent assay (ELISA) test has been developed that is suitable for use in all soliped species.8 It is reported to have high sensitivity and can be used as a field test without specialized equipment. All serological tests may be inaccurate for periods up to 6 weeks following the mallein test.
If pus is available, from either open ulcers or necropsy material, the organism can be cultured or the pus injected intraperitoneally into male guinea pigs to attempt to elicit the Strauss reaction. This is a severe orchitis and inflammation of the scrotal sac but it is not highly specific for B. mallei.
Gene sequencing can be used for rapid identification and differentiation from B. pseudomallei.9
In the acute form there are multiple petechial hemorrhages throughout the body and a severe catarrhal bronchopneumonia with enlargement of the bronchial lymph nodes.
In the more common chronic form, the lesions in the lungs take the form of miliary nodules, similar to those of miliary tuberculosis, scattered throughout the lung tissue. Ulcers are present on the mucosa of the upper respiratory tract, especially the nasal mucosa and to a lesser extent that of the larynx, trachea, and bronchi.10 Nodules and ulcers may be present in the skin and subcutis of the limbs, which may be greatly enlarged. Local lymph nodes receiving drainage from affected parts usually contain foci of pus and the lymphatic vessels have similar lesions. Necrotic foci may also be present in other internal organs. B. mallei, and sometimes Arcanobacterium pyogenes, are isolated from infected tissues, and this is main means of confirmation of diagnosis at necropsy.
In live animals that could be carriers, the complement fixation test is used as the official test in most countries. The mallein test is used in those horses whose sera is anticomplementary.
There is little information on treatment. However, it is unlikely that treatment would be an option in most countries. Penicillin and streptomycin have no detectable effect on the progress of the disease but sodium sulfadiazine has been highly effective in the treatment of experimental glanders and melioidosis in hamsters. Treatment for a period of 20 d was necessary to effect 100% recovery. Combinations of a formalized preparation of B. mallei and sulfadiazine, or mallein and sulfadimidine, are reported to be effective in the treatment of affected horses.
Although clinical and serological recovery from glanders occurs occasionally, recovered animals are not solidly immune and attempts to produce artificial immunity have been uniformly unsuccessful.
Complete quarantine of affected premises is necessary. Clinical cases should be destroyed and the remainder subjected to the mallein test at intervals of 3 weeks until all reactors have been removed. A vigorous disinfection program for food and water troughs and premises generally should be instituted to prevent spread while eradication is being carried out. Restriction of the movement of horses should be instituted and the mallein test carried out in horses which may have had contact with the infected group.11
1 Gody D, et al. J Clin Microbiol. 2003;41:2068.
2 Anuntagool N, Sirisinha S. Microbiol Immunol. 2002;46:143.
3 Derbyshire JB. Can Vet J. 2002;43:722.
4 Lopez J, et al. Microbes Infect. 2003;5:1125.
5 Hagebock JM, et al. J Vet Diag Invest. 1993;5:97.
6 Gangulee PC, et al. Ind Vet J. 1966;43:386.
7 Sen GP, et al. Ind Vet J. 1968;45:286.
8 Verma RD, et al. Vet Microbiol. 1990;25:77.
9 Gee JE, et al. J Clin Microbiol. 2003;41:4647.
Several species of the genus Campylobacter are known to cause disease in farm animals; some are potentially zoonotic and the role of some is uncertain. Campylobacter fetus var. venerealis is the cause of infertility and abortion in cattle and the reader is referred to a textbook in theriogenology for more details. C. fetus subsp. fetus causes sporadic abortion in cattle and enzootic abortion in sheep and has been associated with bacteremia in man.1 The organism has also been isolated from the intestines of healthy sheep and cattle and from enteric lesions in cattle with enteritis, but its significance as the causative agent is uncertain.
Campylobacter jejuni and Campylobacter coli can be isolated from the intestines of healthy farm animals, poultry, pets, zoo animals, and wild birds.2 C. jejuni and C. hyointestinalis can be found in the rumens and intestines of normal adult cattle and calves. C. jejuni and C. coli were isolated from rectal swabs of dairy cows in New Zealand where the prevalence was 24%, 31%, and 12% in the summer, autumn, and winter, respectively. C. jejuni and C. coli were present in the feces of slaughter-age cattle and sheep in Australia with median prevalences and ranges for dairy cattle, 6% (0–24%), feedlot beef cattle, 58% (12–92%), pasture beef cattle, 2% (0–52%) mutton sheep, 0% (0–4%), and prime lambs 8%.3 The cattle production system being used may be an important risk factor. The organism may be present in about 15% of cattle at the time of slaughter. C. jejuni is widely distributed among northwestern US dairy farms, while C. coli is more narrowly distributed, more particularly in calf rearing farms.4
Approximately 60% of the specimens of healthy slaughter pigs may yield C. jejuni.2 C. coli has been isolated from the intestinal contents of 99% of pigs at slaughter. A high prevalence of C. coli in the stomach of pigs at slaughter in France is recorded, and a high proportion of the strains were resistant to tetracyclines and erythromycin.5 C. jejuni, C. coli, C. lari can be isolated from pigs on commercial swine herds.6 Piglets probably become colonized with Campylobacter within a few hours of birth.
The prevalence of Campylobacter infections in both diarrheic and non-diarrheic calves, piglets, lambs, and goat kids may average around 50% but there is no correlation between the occurrence of the organism in the feces and the presence of diarrhea. However, the presence of these organisms constitutes a potential zoonosis among animal handlers. The details of surveys of the incidence of campylobacters from the tissues of cattle at slaughter and from fresh and frozen meat and poultry collected at slaughter are available.3 An adaptation of the ELISA test is available for the detection of antibodies to Campylobacter sp. for use in seroepidemiological studies in herds of cattle and sheep. Wild birds probably constitute the main natural reservoir of infection.
Individual single-visit farm prevalence of intestinal Campylobacter, predominantly C. jejuni, in lactating dairy cows from various regions of the US ranges from 0 to 10%, and there was no difference geographically.7 This low prevalence compares favorably with the rates of 5% for beef cattle on pasture,8 with 7% of dairy cows in the UK,9 and with 6–7% of adult cattle in the US.8 Using a PCR assay of the feces for C. jejuni, 80% of dairy herds were positive, and 38% of individual cows were positive.10 Possible risk factors for C. jejuni were application of manure with broadcast spreaders, feeding of whole cottonseed or hulls, and accessibility of feed to birds.
In feedlot beef cattle, 100% of the animals may shed campylobacters over a period of several months.11 In surveys of feedlot cattle in Ireland over a period of several months, 54% of the animals shed Campylobacter and Campylobacter coli, 69 and 30%, respectively.12 Of environmental pen samples, 29% were positive, and C. jejuni and C. coli accounted for 35 and 59%, respectively. Campylobacter was not isolated from any of the dressed carcasses.
Abortions in beef cattle herds have been attributed to C. jejuni.13 The abortion rates were 19% and 10% in two herds on neighboring ranches in Saskatchewan. Abortions occurred in late gestation and were accompanied by retention of fetal membranes and weight loss. Necrotizing and suppurative placentitis and fetal bronchopneumonia along with culture of large numbers of C. jejuni from the placental and fetal tissues were present. The organism was also isolated from the feces of aborting and healthy cows, and diarrheic and healthy calves. It is suggested that the source and mode of transmission of the organism was fecal contamination of water supplies and feeding grounds by carrier cows or wildlife. C. coli alone and C. jejuni and C. fetus subsp. fetus together have been isolated in epidemics of abortion in sheep. The IV inoculation of C. jejuni into pregnant sheep results in abortion 7–12 days later. A purulent endometritis and vasculitis were present.
C. jejuni has been isolated from an aborted fetus from a goat and the fetus of a heifer which aborted.
Campylobacter jejuni is adapted to the intestinal tract of warm-blooded animals and does not normally replicate outside this environmental niche.14 The single polar flagellum and corkscrew shape facilitates motility in the viscous intestinal mucus. The bacterium gradually dies outside the host’s intestinal tract. C. jejuni strains could not be isolated from water after 3 weeks but may survive for up to 60 days in unstirred water. The distribution and diversity of campylobacters in a large-scale farming environment in the UK was determined by systematic sampling of feces, soil, and water.15 The organism was widespread, there were low levels of antibiotic resistance, high genetic diversity, and a strain of C. coli which may have become adapted to survival or persistence in water.
The organism is not normally pathogenic in farm animals. In humans, the infectious dose is considered to be <1000 Campylobacter organisms.14
Increasing antimicrobial resistance in Campylobacter is being recognized worldwide, and resistance to the quinolones is most common in isolates of both C. jejuni and C. coli from food-producing animals, especially poultry.16,17 Resistance of C. jejuni from poultry increased to 30% within several years after its approval for use as mass water medication in poultry.18 The antimicrobial susceptibilities of Campylobacter spp. isolated from organic and conventional dairy herds in Wisconsin, US, were not different.19 Thus the restricted use of antimicrobials on organic dairy farms had no effect on antimicrobial resistance to ciprofloxacin, gentamicin, erythromycin, and tetracycline in Campylobacter spp. From dairy farms in Washington State, C. coli isolates were more frequently resistant than C. jejuni to ciprofloxacin, nalidixic acid, erythromycin, and doxycycline.4 Of C. coli isolates from dairy calf rearing farms, 89.3% and 72.2% from feedlots were resistant to quinolone antimicrobials, respectively. Multidrug resistance was more common among C. coli than C. jejuni.
Campylobacter is the leading bacterial cause of diarrhea in humans in many industrialized countries. In the United States, disease caused by C. jejuni or C. coli has been estimated to affect 7 million people annually, causing 110–511 deaths.20 Data from population-based studies indicate that the most important cause of indigenous foodborne disease is contaminated chicken.21 Red meat (beef, lamb, and pork) also contribute to illness despite the lower risk.
C. jejuni is ubiquitous in areas contaminated with cattle feces but the intensity of infection varies considerably. Humans can be exposed to Campylobacter spp. in a range of sources via both food and environmental pathways. In dairy farm areas in which there are also outdoor recreational activities, Campylobacter spp. have been isolated from a range of environmental samples by use of a systematic sampling grid.22 C. jejuni was the most prevalent species in all animal species, ranging from 11% in samples from nonavian wildlife to 36% in cattle feces, and from 15% of water samples. C. coli was most commonly found in water (17%) and sheep (21%). C. lari was commonly found in water and in birds. Many of the C. jejuni genotypes isolated from cattle, wildlife, and water were indistinguishable from those recovered from human clinical cases.22
The annual increase in Campylobacter infections in England and Wales begins in early May and reaches a peak in early June. This seasonal incidence may be associated with transmission of the organism by flies.23
An estimated 20% of cases of illness associated with C. jejuni are due to vehicles of infection other than food, including water. Campylobacter spp. have been found to cause waterborne outbreaks worldwide, especially where people drink untreated water from streams and other sources.20 Untreated surface water has been implicated in Campylobacter outbreaks in New Zealand, Finland, England, Wales, Australia, the United States, and Canada. The Walkerton, Ontario, waterborne outbreak of 2000 resulted from entry of Escherichia coli 0157:H7 and Campylobacter spp. from neighboring cattle farms into the town water supply.20 The bacteriologic, epidemiologic, and hydrogeologic data indicated that the bacteria from cattle manure were able to enter groundwater after heavy rains and contaminate a well serving the town of Walkerton, subsequently infecting those consuming the water. Some investigations consider that beef cattle represent a limited threat to water supplies and subsequent transmission of Campylobacter to humans.24
Raw milk contaminated by infected cows is a major cause of food-borne human campylobacteriosis in the United States and United Kingdom.14 C. jejuni, along with other food-borne pathogens such as shiga-toxin producing E. coli, Listeria monocytogenes, Salmonella spp., and Yersinia enterocolitica can be found in the bulk tank milk supply of dairy herds in Canada and the United States.25 Drinking raw cows’ milk is commonly related to illness due to C. jejuni.2 Raw goats’ milk may transmit C. jejuni infection from animals to humans.2 There is a strong association of Campylobacter infection in humans with residence on a farm14 and contact with diarrheic animals is a major risk for Campylobacter enteritis in humans. Fecal contamination, rather than udder infection, is considered to be the means by which campylobacters enter milk and thereby infect humans. C. jejuni has been isolated from the bulk milk supply of goats whose milk was associated with Campylobacter infection in a human.
Along with other food-borne bacteria, such as E. coli, Salmonella spp., Campylobacter spp. can be found in retail raw meats (chicken, turkey, pork, and beef) sampled in supermarket chain stores.26
There is good evidence that isolates of C. jejuni from human disease and farm animals are very similar. The use of multilocus sequence typing is being used to compare the genotypes of C. jejuni from farm animals and the environment with those from retail food and human disease.15,27
The role of C. jejuni as primary pathogens in farm animals is uncertain. The organism was originally thought to be the causative agent of winter dysentery in cattle but reliable evidence for this relationship has not been found. Experimentally, the organism will cause a mucoid diarrhea, often with dysentery and a fever in calves.
C. jejuni or C. coli can cause a mild self-limiting enteritis and bacteremia when inoculated orally into newborn calves. The organism has been isolated from the feces of diarrheic calves and lambs, which suggests that it may be a causative agent in some outbreaks of diarrhea but this has not yet been substantiated. The oral inoculation of pure cultures of C. fetus subsp. intestinalis into young calves will also result in an enteritis similar to that associated with C. jejuni. The oral inoculation of C. jejuni into gnotobiotic pigs results in diffuse edema and neutrophil infiltration of the mucosa of the cecum and colon. An episode of diarrhea in calves 5–12 weeks of age has been attributed to C. hyointestinalis. A Campylobacter-like organism has been isolated from young sheep about 1–2 months after weaning, when they were about 6 months of age, affected with weaner colitis. The morbidity rates in flocks ranged from 20–75% and the case–fatality rate was about 3%. Campylobacter spp. can be found in a high proportion of foals on horse farms where persistent non-responsive diarrhea has been a problem. Outbreaks of severe gastroenteritis in fattening lambs have been attributed to C. jejuni, these outbreaks were treated successfully with daily injections of erythromycin followed by a single injection of long-acting oxytetracycline.
Campylobacter fetus subsp. fecalis has been isolated from intestinal lesions of cattle and experimentally will cause a diarrhea and dysentery in calves.
Campylobacter coli (formerly Vibrio coli) has been isolated from the small intestines of diarrheic piglets and experimentally can cause colitis in young piglets. The organism may be the cause of naturally occurring diarrhea in nursing piglets and weaned pigs in certain circumstances.
The disease may be so mild as to be unapparent, without fever, and may be manifested only by mild depression and soft feces with occasional strands of mucus.
The information on the various methods used for the detection and identification of Campylobacter in laboratory samples has been reviewed.28 Because of the unique growth characteristics of Campylobacter, isolation of these organisms from field samples requires the use of special media and culture conditions, and is generally laborious and time-consuming. However, isolation of Campylobacter from feces is possible with high success rates. Recovery of Campylobacter from environmental samples can be difficult because the organism does not propagate in the environment.28 The use of molecular detection methods has greatly facilitated the specific and rapid detection and identification of campylobacters, but has not replaced the gold standard of traditional culture methods. Detection and quantification of C. jejuni in the feces of naturally infected cattle is possible using real-time quantitative PCR.29
The laboratory methods used to distinguish epidemic-associated Campylobacter strains isolated from animals and humans have been examined.7
At necropsy, there may be a diffuse catarrhal to severe hemorrhagic enteritis of the jejunum and ileum.
Control depends on sanitation and hygiene in livestock barns to reduce the bacterial populations in the environment of the animals. The numbers of organisms can be reduced and controlled in meat processing plants by using Hazard Analysis of Critical Control Points including the washing, handling and freezing of carcasses. Improvement of food-handling skills in restaurants and in the home kitchen will reduce transmission of the organism and adequate cooking of raw meat such as poultry to an internal temperature of 82°C will eliminate the organism.14
1 Delong WJ, et al. Am J Vet Res. 1996;57:163.
2 Altekruse SF, et al. J Am Vet Med Assoc. 1994;204:57.
3 Bailey GD, et al. Commun Dis Intell. 2003;27:249.
4 Bae W, et al. Appl Environ Microbiol. 2005;71:169.
5 Payot S, et al. Vet Microbiol. 2004;101:91.
6 Young CR, et al. Res Vet Sci. 2000;68:75.
7 Harvey RB, et al. J Food Prot. 2004;67:1476.
8 Beach JC, et al. J Food Prot. 2002;65:1687.
9 Ataby HI, et al. J Appl Microbiol. 1998;84:733.
10 Wesley IV, et al. Appl Environ Microbiol. 2000;66:1994.
11 Inglis GD, et al. J Appl Microbiol. 2004;97:410.
12 Minhan D, et al. J Vet Med B. 2004;51:28.
13 Van Donkersgoed J, et al. Can Vet J. 1990;31:373.
14 Altekruse SF, et al. J Am Vet Med Assoc. 2003;223:445.
15 Leatherbarrow AJH, et al. Appl Environ Microbiol. 2004;70:822.
16 Pezzoti G, et al. Int J Food Microbiol. 2003;82:281.
17 Ishihara K, et al. Int J Antimicrobiol Agents. 2004;24:63.
18 Lucey B, et al. Vet Rec. 2002;151:317.
19 Sato K, et al. Appl Environ Microbiol 70:1442.
20 Clark CG, et al. Emerg Infect Dis. 2004;9:1232.
21 Adak GK, et al. Emerg Infect Dis. 2005;11:365.
22 Brown PE, et al. Appl Environ Microbiol. 2004;70:6501.
23 Nichols GL. Emerg Infect Dis. 2005;11:361.
24 Hoar BR, et al. Epidemiol Infect. 2001;127:147.
25 Jayarao BM, Henning DR. J Dairy Sci. 2001;84:2157.
26 Ohya T, et al. Vet Rec. 1999;145:316.
27 Colles FM, et al. Appl Environ Microbiol. 2003;69:7409.
28 Sahin O, et al. Torrence ME, Isaacson RE, editors. Microbial food safety in animal agriculture. Iowa State University Press. 2003:183.
29 Inglis GD, Kalischuk LD. Appl Environ Microbiol. 2004;70:2296.
Four to 8 weeks after weaning, feeder pigs, and young gilts, sows and boars. Risk factors not known
Diarrhea, weight loss, inappetence, and may recover. Outbreaks of bloody diarrhea and rapid death may occur in feeder pigs, young gilts, and boars
The causative agent is Lawsonia intracellularis (LI), which was isolated in 1993.1 It was described about 60 years earlier and first reported in 1974.2 Koch’s postulates for the disease were fulfilled in 1993.3 It is an obligate intracellular bacterium4 or in other words host cell dependent. It was also known as ileal symbiont intracellularis. Recent classification of DNA from infected enterocytes indicates a close relationship to Desulfovibrio species.4 It has also been shown to be closely related to Bilophila wadsworthii which is a known inhabitant of the human colon and may be associated with appendicitis and is widely found in pigs in Australia.5 Molecular typing of the organism has recently been described.6 The isolates of Lawsonia intracellularis from the United States are similar to the European isolates.7
Formerly, the intracellular organisms, Campylobacter sputorum subsp. mucosalis and Campylobacter hyointestinalis were considered to be the causative agents because they could be isolated from the intestines of pigs with proliferative ileitis. Both pure cultures and mucosal homogenates of LI will produce clinical signs, lesions, and shedding.
The disease complex, often just called ileitis, occurs in two forms; a chronic form usually referred to as porcine intestinal adenomatosis (PIA) or necrotic enteritis occurs from 6–20 weeks and an acute form called porcine hemorrhagic enteropathy or regional ileitis occurs earlier from 4–12 weeks.
The worldwide occurrence has been described.8
The porcine proliferative enteropathy complex affects pigs from weaning age to feeder pigs and also young gilts, sows, and boars. It is characterized clinically by diarrhea, loss of body weight and inappetence in recently weaned pigs, and sudden death in feeder pigs, young gilts, and boars. The essential lesions are proliferative and there seems to be an etiological and pathological relationship between porcine intestinal adenomatosis, necrotic enteritis, regional enteritis, and hemorrhagic enteropathy. A study in Belgium suggested that 24% of slaughtered pigs had a thickened ileum with a range in farm batches from 10–49%.9 In Denmark 94% of herds were infected with a mean within herd prevalence of 30%.10 In Canada, there is a widespread distribution between 50–100% of herds in the provinces with 5–89% of pigs affected.11 In the USA it was found using the IPMA test to study antibodies that 75% of growing herds had antibodies and within the herd the prevalence was 11–91%. Of the breeding herds 78% had antibodies with two peaks at the time of infection and 9–18 weeks later12 and with an overall prevalence of 5–61%.
In Canada, studying 96 cases of porcine proliferative enteropathy,13 it was found that 15% were in weaners (8–10 weeks), 36% in growers from 10–18 weeks and 14% amongst finishers of 18–26 weeks. A further 16% were in mature pigs of >26 weeks.
Estimation of the incidence of disease is complicated by the difficulties in making an accurate clinical and pathologic diagnosis. Surveys of pig farms in Australia indicate that 56% had either observed the disease or the veterinarian had made the diagnosis.14 Non-hemorrhagic proliferative enteritis occurred most commonly in pigs 6–24 weeks of age. Proliferative hemorrhagic enteropathy usually affects pigs over 16 weeks of age but occurs in pigs as young as 6 weeks and as old as 4 years of age.14
Proliferative hemorrhagic enteropathy is one form of the proliferative enteropathy complex and has been reported from the United Kingdom, Europe, Australia, Asia, and the United States, and appears to occur in most countries. It probably has a worldwide distribution8 with 30–60% of herds affected depending on the country. In Germany 82.7% of finishing herds had seroconversion.15 It is especially common in hysterectomy-derived or specific pathogen-free (SPF) herds and has a higher prevalence in the hot summer period. In some countries its prevalence is increasing and it is emerging as a major syndrome in SPF herds.
The disease in all ages is frequently associated with the concurrent occurrence of porcine intestinal adenomatosis, but it is unknown whether the hemorrhagic syndrome results from some insult to the intestine which also predisposes to a proliferative enteropathy or whether it is simply an acute manifestation of this disease. The related syndromes of necrotic enteritis and regional ileitis can be found in apparently healthy pigs examined at slaughter. Because the disease is common in pigs, suboptimal growth of pigs in nutritional studies may be due to the disease complex.16 It has been suggested that it can live extracellularly within the environment for two weeks17 at 5–15°C. It appears highly resistant to a lot of cleaning agents such as pevione–iodine, or K permangosulfate but may be susceptible to 3% cetrimide. In one study transmission occurred despite cleaning, use of footbaths, and dedicated boots, etc.18
It has been suggested that it normally lives in organic matter in weaner units awaiting the arrival of batches of susceptible pigs with the resultant sudden increase in shedding 4–12 weeks after weaning.19 The recent finding of LI in the tonsil may be a coincidental finding in that they have just been trapped in the crypts after licking of infected material20 as they were only found in this site in 2/32 pigs. Mixed infections are found in 10% of growers and there is a strong association between diarrhea and prevalence of B. hyodysenteriae and B. pilosicoli.21
The potential economic effect can be quite severe with estimates of $1.48–3.42 for mild infections and $11.50–22.19 for severe infections.22
Surveys of fecal samples from swine herds in Taiwan revealed an overall prevalence of infection of Lawsonia intracellularis in 30% of herds and 5.5% of pigs.23 They have looked for it in wild pigs in Sweden but not found it.24
The disease can occur in all ages of postweaned pigs, but it has a high incidence in young replacement gilts and boars at 6–9 months of age and in pigs approximately 4–8 weeks after weaning. The high incidence in replacement gilts may be due to suppression of the disease by low-level feeding of antibacterial agents during the growing period, but frequently the syndrome appears first in gilts and some time later in the growing pigs. In gilts, outbreaks may be explosive, but generally are short-lived with morbidity rates of up to 50% of the group occurring within a 2- to 3-week period. The case–fatality rate does not usually exceed 10%. In large herds with continual addition to the replacement gilt herd and in herds where the disease occurs in grower pigs, outbreaks may be more prolonged. The disease in growers generally has equivalent morbidity and case–fatality rates, but is more severe in that runting of surviving and contemporary pigs may occur necessitating further economic loss through culling.
When given experimentally at a high level of 109–1010 LI per pig mortality in the untreated groups varied from 10–50% which is considered much higher than in the natural outbreak.25,26
There may be two patterns of infection. One is an early infection and the second is a delayed infection which is seen in farms which have separation of pigs at weaning and all in/all out methods of production.8
Very little is known about the risk factors. A gene has been discovered which encodes for a surface antigen (LsaA) which is believed to be associated with attachment to and entry into cells and which is synthesized during infections.27 A study of recorded outbreaks of proliferative hemorrhagic enteropathy indicated the disease often occurred within 12 months after repopulation of the herd, and following withdrawal of antimicrobials from the feed.28 It has been proposed that the introduction of breeding stock from herds where the disease is endemic may be a risk factor but this is not documented. In a study in the UK29 there seemed to be a higher risk where there were over 500 sows. An older parity structure in the sow population seemed to reduce infection. There seemed to be a higher risk if buying in boars. Fully slatted or fully meshed floors also carried a higher risk of infection compared with solid floors or straw. A higher risk was seen in those herds where large numbers of pigs entered the finishing units simultaneously. Pigs on concrete slats may also be predisposed. Intensive systems were more severely affected than outdoor systems. There was a reduced risk if there was thorough cleaning and disinfection (all in/all out) before the next group of pigs arrived.30 Seroconversion usually occurred as the pigs entered the finishing site suggesting that the exposure takes place in the nursery.31 A recent study in the USA32 identified five major types of risk factor. These were co-mingling, temperature fluctuations (overheating/chilling), transportation, depopulation and new buildings. Sows may have low levels of antibody and are capable of passing on colostral protection to the piglets. Maternal antibodies have usually declined by 3–5 weeks of age but may be extended to 42 days by repeated sow vaccination33 by which time exposure may have occurred and therefore there may be both active and passive antibodies.31
The organism is found in hamsters, ferrets, foxes, hares, deer, emus, ostriches, and primates. The significance of these alternative hosts has not yet been ascertained. Most significant host is the pig. The organism can be spread by both growing pigs and adults. Gilts can be shedders and carriers. It can probably survive in the extra-cellular world for 1–2 weeks at 5–15°C.17
The method of transmission between pigs is assumed to be the fecal–oral route but no information is available to support this.
The infection process appears to go ileum to colon, to cecum and finally rectum. The histological lesions may have cleared from the ileum by day 29 following inoculation.34
Proliferative enteropathy is characterized by the hyperplasia of the epithelial cells of the intestinal crypts, particularly in the ileum and colon. The presence of non-membrane bound, curved bacteria free in the cytoplasm of the affected enterocytes is a consistent feature of the disease.35 These bacteria have been cultured in a rat enterocyte cell line36 and the disease has been reproduced in hamsters by inoculation of a pure culture of the organism derived from pigs.37 The disease has been reproduced experimentally by inoculation of conventional pigs with the organisms.38,39 The organisms infect the immature cells of the mucosal glands and stop them from maturing. This causes them to multiply without leaving the gland and they then degenerate and the glands continue to proliferate. Gross and microscopic lesions typical of acute proliferative enteritis can be reproduced by inoculation of cell-cultured Lawsonia intracellularis into pigs 3 or 7 weeks of age.39 The incubation period is about 7–14 days with the early lesions appearing in the terminal ileum. Fecal shedding usually occurs about 7 days post-challenge and the animals seroconvert about 14 days post-challenge. The disease peak is about 21 days post-infection. The clinical signs decrease and the lesions begin to resolve after 28 days. The disease process results in a 2 week delay in marketing. Inoculation of gnotobiotic pigs does not cause the disease. It now seems certain that Lawsonia intracellularis is the causative agent of the disease complex.40 Infection of intestinal epithelial cells is causally linked to marked hyperplastic proliferation of affected tissue.41Lawsonia intracellularis is an obligate intracellular bacterium which causes hyperplasia of intestinal tissues; this eventually reverts to normal. The organism internalizes and multiplies within the cells and it is proposed that the organism is capable of affecting, directly or indirectly, the cell cycle within the intestinal epithelium. This may or may not be concerned with the role of cyclin kinase p27 which regulates differentiation of immature crypt cells into the differentiated form.42 The changes in the experimental disease are similar to those in the natural disease. Following experimental infection there is almost complete replacement of normal ileal mucosa by adenomatous mucosa. Affected crypts are enlarged and branched, with loss of goblet cells and marked proliferation of crypt epithelial cells. Hyperplastic lesions may develop 2–3 weeks after challenge and persist for several weeks. In older animals, the lesions may be complicated by acute mucosal hemorrhage or necrosis. In the progressive stage of the disease, 3 weeks after infection, numerous organisms are consistently present within affected intestinal epithelial cells but not elsewhere.41 In the developed and recovering stage of the disease, 7–9 weeks after infection, ultrastructural features in affected intestinal tissues consist of pale, swollen, protruding epithelial cells and shrunken epithelial cells. This is followed by the appearance of apoptotic bodies in both epithelial cells and macrophages, the reappearance of normal goblet cells and reduced numbers of organisms within the lesions. Bacteria are released from cells via cytoplasmic and cellular protrusions into the intestinal lumen and can be found in fecal samples.
In the experimental disease in pigs, seroconversion to the organism does not occur, confirming the weak response characteristic of the natural disease.43
The proliferative lesion may result in suboptimal performance in otherwise normal pigs or unthriftiness, or be manifested as acute intestinal hemorrhage during the recovery stages of intestinal adenomatosis. The hemorrhagic lesions are more difficult to explain but there may be direct or indirect toxic damage to the endothelium of the blood vessels.
It has been suggested that there is a close association between the presence of LI and reduced T- and B-cell numbers. This provides evidence of an immunosuppressive effect operating in this disease. It seems also that macrophages have an important function with activated macrophages accumulating in the infected hyperplastic glands.44 At day 14 post-infection there were a few pinpoint lesions and the percentage of infected crypts was minimal but at the same time the number of CD3+ cells was reduced and the number of intra-epithelial CD3+ cells was also reduced whilst the CD8 and CD4 cells showed no changes. Apparently there is an induction of an immunosuppressive phenotype with down regulation of an adaptive immune response through the reduction in the CD8+ T- and B-cells.
This disease is one of the common causes of failure to grow, weight variation in batches of pigs and delay to market. Pigs may appear gaunt and may pass watery stools.
Regional ileitis is the most common differential diagnosis of the granulomatous enteritis that is seen in PCV2-associated enteric disease. In many cases both PCV2 and LI have been seen in the same case45 as both target the ileum.46
Porcine proliferative enteropathy (PPE) or ileitis occurs in pigs 6–16 weeks of age. In the chronic form, a reduction in growth rate and failure to thrive are common. Affected pigs are afebrile and diarrhea occurs, but is unremarkable. Most cases recover spontaneously within 6 weeks of the onset of signs. When inflammation and necrosis have resulted in necrotic enteritis and regional ileitis, diarrhea and severe weight loss occur followed by death, often by ileal perforation in the case of regional ileitis.
Proliferative hemorrhagic enteropathy (PHE) occurs in older pigs such as young gilts and boars and is manifested primarily by bloody diarrhea and sudden death. Others within the group may show skin pallor and hemorrhagic feces with fibrin casts but otherwise appear clinically normal. In some pigs there is continual blood loss and death occurs within 48 h of the onset of hemorrhage, but in the majority the hemorrhage is transient. In outbreaks, up to 70% of pigs affected with dysentery may die within 24 h after the onset of signs.28 Fever is not a feature and the majority of pigs suffer only a minor setback for a 2-week period. A small percentage develop chronic illthrift.
In grower pigs the disease is economically more severe. As in gilts, acute death with marked skin pallor and without premonitory signs can occur but survivors show illthrift and as the outbreak progresses contemporary pigs may show a chronic syndrome of illthrift with the periodic passage of bloody feces.
The most staggering advance in clinical diagnosis may be by use of ultrasonics,47 where it is said that it is possible to diagnose the condition by measuring the thickness of the ileal mucosa. It was said to increase from 0.27–0.36 cm in normal animals to 0.30–0.70 in affected animals.
The organism can be detected in the feces of healthy 10- to 25-week-old growing/finishing pigs, which is probably the age group of pigs serving as the main source of infection for younger nursery pigs.
A polymerase chain reaction assay is highly reliable for the detection of the organism in feces and intestinal tissues.48,49 It may detect as few as 2 × 102 bacterial cells per gram of feces50 but more likely is that the PCR detects shedding of 103 or greater per gram of feces.51,52
Positive results with the PCR are only present in animals with active lesions of proliferative enteropathy.53 Shedding as detected by PCR may start as early as 6–8 weeks and continue to 28 weeks. From seroconversion to first shedding was 2–8 weeks.
A fluorescent in situ hybridization technique targeting 16S ribosomal RNA using an oligonucleotide probe successfully identified LI.54
Seroconversion may commence between 12–27 weeks. The range for positivity from first detection was 7–23 weeks.
The immediate impact is of a thickened ileum and cecum and less frequently a spiral colon. Not all cases have lesions. Some may be so mild as to be overlooked. Obvious gross lesions occur in severe cases but in the less severe, histology is needed. The pathology is related to the dose.55 As long as you remember these facts you can monitor LI in the abattoir.56
A complex gross, histological and immunohistochemical study of LI has been made57 in which the pigs showed complete recovery and were IHC –ve by 35 days post-infection. The antigen was detected in the intestine, lymph node (macrophages) and in the tonsils (free living in the crypts). They were found in the rectum and in several portions of the large intestine. The first site of colonization was the jejunum and ileum and then the lower intestinal segments. On day 29 there was nothing in the small intestine but the LI were still observed in the cecum, proximal colon, and rectum. Mucosal IgA was first detected on day 15 and was still detectable on day 29 but in all cases the titers varied from only 1:4–1:16.
The macroscopic lesions of proliferative enteropathy were first detected at 11 days post-infection which is the same time as histological identification with enterocyte hyperplasia and reduced goblet cells. Immunohistochemical identification can be seen at 5 days post-infection and continues until day 29.
In porcine intestinal adenomatosis, the prominent lesions are in the terminal ileum and proximal portion of the large intestine. There is gross thickening of the mucosa and submucosa of the terminal ileum and the colonic mucosa may also appear congested and slightly thickened.
In both forms of the disease the mucosal surface may be eroded and may look granular with abundant adherent material in the form of fibrinonecrotic debris. There may also be a fibrinonecrotic core filling the lumen. In PHE the only difference may be that the surface of the mucosa may be covered by large undigested blood clots.
Histologically, the mucosal change consists of marked proliferation of immature epithelial cells and a suppurative cryptitis.
In regional ileitis (called hosepipe gut) the distal ileum is rigid due to thickening of the intestinal wall caused by muscular hypertrophy and granulation tissue formation. The initiating mucosal damage is often somewhat masked due to colonization of the ulcerated mucosa by secondary bacterial invaders.
In proliferative hemorrhagic enteropathy, the carcass is usually very pale and massive amounts of blood are often present within the intestinal tract. The mucosa and submucosa of the ileum are thickened and may be coated in fibrin. Fibrin casts are also sometimes present. Although the intestinal wall is dark red and hemorrhagic, there may be no obvious points of hemorrhage. Histologically, there is evidence of vascular congestion, fibrin thrombi, increased permeability of blood vessels and necrosis of the intestinal mucosa. The character of the vascular lesion resembles an acute bacterial infection and type I hypersensitivity reaction. Again, the key microscopic feature is the presence of proliferating immature epithelial cells with basophilic nuclei, which line the greatly elongated crypts. There are no goblet cells in this site. In an analysis of histological lesions crypt abscesses were seen in 20% of pigs, decreased goblet cells in 90%, hypertrophy and hyperplasia in 3%, hypertrophy of both muscle coats in 78%, increased eosinophils in 34% and lymphoid hyperplasia in 90%.
In chronic cases the lesions described above are nearly all replaced by fibrous connective tissue and the diagnosis may rely on seeing just isolated pieces of mucosa.
Lawsoniana are also a common cause of colitis. In 70% of cases of colitis LI are also found in the colonic mucosa. In three cases, LI were found only in the colon and in these infected large bowels there was an excess of mucus on the surface.58
Staining of smears of ileal mucosa with modified acid-fast stains may reveal typical curved bacterial rods in the apical cytoplasm of the infected proliferating enterocytes, permitting a presumptive diagnosis. It is not always specific for Lawsoniana. They are not always present in necrotic debris or autolysed tissue. Immunohistochemistry or silver stains (Warthin/Stary)59 of formalin-fixed gut are usually sufficient to detect the intracellular organisms in all forms of proliferative enteritis. Lawsonia intracellularis can also be identified using a PCR assay.49,53 It is possible to find bacterial antigen in the lamina propria and draining lymph nodes of the ileum and this is a result of the natural process of infection clearance.57
• Bacteriology – distal ileum, proximal colon (DIRECT SMEAR, PCR). The organism needs to grow on tissue cell lines at oxygen and CO2 concentrations that mimic the small intestine. It is not really an option because these techniques are difficult and the organism is an obligate intracellular organism.
There has been a considerable development of PCR techniques for feces as an antemortem technique.60,61 This is a variable sensitivity which is affected by sample quality and the presence of inhibitory factors in feces, but the specificity is around 97%. It appears to be very useful in the clinically ill but not so reliable in the subclinically affected. The PCR is more specific when applied to the ileal mucosa rather than to feces. It has been reported that fecal samples are more likely to be PCR positive in herds with PHE rather than in PIA herds. It is more sensitive than either WS staining or IFAT.62 Shedding commences around 7 weeks and is observed most between 13–16 weeks.63,64 A one tube nested-PCR has been developed which is very sensitive and less prone to false positives compared to a standard nested-PCR.65 A 5′ nuclease assay has been developed with a detection limit of 1 LI cell per PCR tube.66 A real-time PCR has been designed67 as a high-throughput test for use on feces. It is as specific as a conventional PCR but is more sensitive. It can be quantified and can be carried out with pure cultures, tissue homogenate or bacteria shed in feces.
A multiplex PCR has also been described for B. hyodysenteriae, pilosicoli, and L. intracellularis. It has a 100% specificity for the three species and does not generate false positives.68
There is also an indirect fluorescent technique but this requires expertise and a reliable Lawsoniana specific antibody and again is not 100% for subclinically affected animals.69,70 The percentage of agreement between IFAT and IPMA was 98.6%.71 It has been suggested that IFAT is more sensitive than PCR in antemortem testing.50
• Histology – distal ileum, proximal colon (LM, IHC). Immuno-histochemistry was described72-75
• Serology – current methods utilize LI grown in enterocytes or LI prepared on slides as the antigen. These assays are specific because cell cultures or slides are examined microscopically and specifically stained bacteria can be distinguished from any background. Staining of bacteria is either by a fluorescent (IFA) or peroxidase-labeled IPMA. The IPMA test is highly specific (100%) and fairly sensitive (90%) in experimentally infected animals.71,76 It is an appropriate diagnostic test for herd screening but not for diagnosing PPE on an individual animal basis. The IgG antibodies may be only short lived and found only between 18–24 weeks. These have proved useful for routine PPE diagnosis, although the humoral response is often weak and short lived. Titers of 1:30 to LI appear about 2 weeks after infection and 90% become positive by about 3 weeks after challenge with 5% having titers of 1:480 or above. They are however already decaying by about 4 weeks after challenge. Antibody was not detected until 16 weeks of age and often not until 19–22 weeks71
• A cell-mediated response can be detected in the research laboratory using an (enzyme linked immunospot assay) Elispot-T-cell assay that measures the LI specific secretion of IFN-γ by lymphocytes. It appears to follow the same pattern as the humoral response and it also starts to decay from about 3 weeks although more slowly.
Both humoral and cell-mediated responses can still be detected 13 weeks after challenge or vaccination.71,76
Characteristic clinical findings are inappetence, loss of weight and mild diarrhea in recently weaned pigs. Must be differentiated from postweaning coliform gastroenteritis – clinically much more severe and death rapidly occurs. The postweaning drop in average daily gain (postweaning check) occurs within several days after weaning and recovery occurs within several days following consumption of a normal daily intake of feed.
Occurs in feeder pigs, young gilts, and boars and is characterized by sudden death and extreme pallor of the skin. Must be differentiated from fatal hemorrhagic esophagogastric ulceration, acute swine dysentery, and intestinal hemorrhage syndrome.
Occurs in all ages of pigs but especially in growers. The necropsy finding of ulceration in the non-glandular portion of the stomach at the esophageal entrance along with hemorrhage into the stomach with passage into the intestines provides easy differentiation. Acute death with intestinal hemorrhage occurs occasionally in swine dysentery. More common in adults affected with the disease and at the onset of an outbreak, skin pallor is not as marked and hemorrhage is restricted to the large intestine, and is associated with the characteristic lesions of swine dysentery in this area. Contemporary pigs show clinical and necropsy findings typical for this disease and the diagnosis can be confirmed with laboratory studies.
More difficult to differentiate from the proliferative hemorrhagic enteropathy. Occurs most commonly in 3–6-month-old pigs that are well-nourished and many but not all outbreaks have been associated with whey feeding. Typically associated with abdominal distension and evidence of abdominal pain preceding death and the presence of marked intestinal tympany on postmortem examination. In many cases, hemorrhage in the intestine appears to result from torsion which occludes the mesenteric veins. It occurs in all areas of the intestine except the proximal duodenum and stomach, which have separate drainage. Owing to intestinal distension the torsion may be easily missed but it is best determined by the abnormal cranial direction of the blind end of the cecum and palpation of the mesentery. This distribution of hemorrhage may occur without the occurrence of torsion and the etiology in these cases is unknown.
Biosecurity to prevent the entry of infection is the key to control. Beware of carrier pigs, isolate for 30–60 days, use preventive antibiotics as outlined below, use laboratory diagnostics and vaccinate using the new water vaccine.
Eradication using early weaning is not a possibility, but using medication and vaccination is a possibility.
It has been said77 that pigs between 30–50 kg shed fewer LI in the feces when they are fed non-pelleted and non-heated (home-mixed) feed.
An eradication scheme for LI used in Denmark following the use of antimicrobials (tiamulin, lincomycin, and tylosin) failed.78
A control program was tried in the UK using PCR to identify affected animals and medication with chlortetracycline and tiamulin for control.79 The number of PCR-positive animals declined from 50–70% to 0%. In pigs over 14 weeks there were some PCR positives derived from treated groups. Another farm used tylosin phosphate and these remained clean.
In acute disease, water medication and particularly individual medications are more effective than treatment through in-feed medication.80
Continuous medication for LI can prevent infection but is frowned upon because it can prevent the development of immunity and extend susceptibility to infection. In fact the timing of any medication can affect the immune response, subsequent fecal shedding and the development of lesions.
There is no published information available on the treatment of individually affected pigs. The disease is usually treated on a herd basis by medication of the feed.
There appears to have been no changes in the in vitro MICs since the 1980s/1990s. There are probably four reasons why medication does not work: (i) underdosing; (ii) concurrent infections; (iii) some other disease or nutrition problem, i.e. misdiagnosis; (iv) antibiotics given too late to be effective.
If you are going to use antimicrobials it is a good idea to start at least 3 weeks before the anticipated acquisition of the infection.
The antimicrobial susceptibility of the organism isolated from pigs with proliferative enteropathy was determined in a tissue culture system. Penicillin, erythromycin, difloxacin, virginiamycin, and chlortetracycline were the most active compounds tested.81 Tiamulin and tilmicosin were the next most active, and the aminoglycosides had the highest minimum inhibitory concentrations. Both lincomycin and tylosin were relatively inactive against the strains of the organism tested.
In the field bacitracin, virginiamycin and salinomycin are useless as are penicillins and fluoroquinolones.
Chlortetracycine, one of the oldest drugs is still used; at 300 or 600 mg/kg it can prevent challenged pigs from developing clinical disease. Chlortetracycline at 300 ppm and tylosin at 600 ppm have prevented the clinical signs of PE.82,83
Tylosin is ideal for treatment by injection, in-feed or through the water and was successfully used for treating PPE at 100 ppm.80 For effectiveness, the antimicrobial would have to accumulate in the cytoplasm of the intestinal cell and block bacterial protein synthesis. The macrolides, tetracyclines, and virginiamycin act by selectively blocking protein synthesis in ribosomes. The oral administration of tylosin phosphate at a dose of 100 or 40 ppm in the feed to pigs for 4 days before experimental challenge and continued for 16 days when the dose was reduced to 40 and 20 ppm was effective in preventing the clinical signs and lesions of proliferative enteropathy.81 It does not appear to block the pattern of seroconversion to LI. Tylosin at 110 ppm significantly reduced fecal shedding of LI and histologic lesions consistent with PPE.84 Injection of tylosin produced an improved diarrhea score, and clinical impression score and thereby weight gain.85 Tylosin tartrate in drinking water for the treatment of ileitis was effective in reducing clinical signs, lesions and reduction in growth rate.86
Lincomycin is ideal for injection, water treatment, and in-feed treatment. Lincospectin at 80 ppm used consecutively was shown to be useful for treatment of PPE.87,88 Lincomycin at 44 and 110 ppm for 21 consecutive days89 was effective in controlling the clinical signs of PPE and at 110 ppm also reduced the mortality associated with PPE. Lincomycin water soluble powder at 250 mL/gal is also effective.90
Aivlosin was found to be useful at concentrations 25% less than those used for tylosin.91
Valnemulin was also shown to be effective at 75 ppm in the feed.92,93
Tiamulin is useful for in-feed medication and water administration. Tiamulin given 50 ppm, 2 days before experimental challenge and kept for 3 weeks prevented the clinical disease.94 In addition, pigs given 150 ppm tiamulin 7 days after challenge remained clinically normal and had no specific lesions of proliferative enteropathy at necropsy. Tiamulin in water is very useful95 but a study showed that in water it interfered with seroconversion whereas administration in feed did not.96
The use of zinc-bacitracin in the feed of growing/finishing pigs at 300 or 200 ppm from weaning up to 100 d of age; or 200 or 100 ppm from 100–125 days of age; and 100 or 50 ppm from 125–156 days of age was effective in controlling the effects of proliferative enteropathy in pigs on a farm with a previous history of the disease.97
Carbadox might have some effect against LI as might zinc oxide.98 It has been shown to be useful if fed in the final 2 weeks in the nursery. It reduces fecal shedding, clinical signs and no IHC+ve or PCR+ve animals were found in one study.99
Hyperimmune chicken eggs for controlling LI infection in growing swine has been described.100
The main difference between respiratory and alimentary diseases in the last few years has been the development of vaccines for the former but not the latter. The recent development of an ileitis vaccine is the first of these for the enteric diseases.
It is a safe, labor-saving, efficient, and easy method of vaccinating pigs through the administration of drinking water using the water proportioner. In the presence of feed medication, vaccinated pigs performed better than the non-vaccinated pigs when exposed to LI challenge. The percentage morbidity was reduced, the feed conversion better and the average daily gain increased by about 6%. There was also a 23% reduction in culls. It is best given in a 7-day antibiotic-free period. The present vaccine is given in water to 70–90 lb gilts. It can be dispensed with antimicrobials and produce protective immunity.101 There is a reduction in gross and microscopic lesions in the complete absence of antimicrobials when the gilts are vaccinated as finishers and the animals receive a booster vaccination every 6 months.102
1 Lawson GHK, et al. J Clin Microbiol. 1993;31:1136.
2 Rowland AC, Lawson GHK. Res Vet Sci. 1974;17:323.
3 McOrist S, et al. Inf Immun. 1995;61:4286.
4 Gebhart CJ, et al. Int J Syst Bacteriol. 1993;43:533.
5 McOrist S, et al. J Clin Microbiol. 2001;39:1577.
6 Beckler DC, et al. Proc 18th Int Pig Vet Soc Cong 2004; p. 249.
7 Knittel JP, et al. Swine Health Prod. 1996;4:119.
8 McOrist S, et al. Pig J. 2003;51:26.
9 Maes D, et al. Vlaams Diergeneesk Tijdschr. 1999;68:231.
10 Stege H, et al. Prev Vet Med. 2000;46:279.
11 Paradis MA, et al. Proc 18th Int Pig Vet Soc Cong 2004; p. 302.
12 Marsteller TA, et al. Proc Am Ass Swine Vet. 2002;33:85.
13 Wilson JB, et al. Can Vet J. 2002;43:604.
14 Holyoake PK, et al. Aust Vet J. 1994;71:418.
15 Wendt M. Proc 18th Int Pig Vet Soc Cong 2004; p. 252.
16 Gogolewski RP, et al. Aust Vet J. 1991;68:406.
17 Collins A, et al. Swine Hlth Prod. 2000;8:211.
18 Moller K, et al. Vet Micro. 1998;62:59.
19 Vestergaard K. Proc 18th Int Pig Vet Soc Cong 2004; p. 831.
20 Jenson TK, et al. Res Vet Sci. 2000;68:23.
21 Lofstedt M, et al. Proc 18th Int Pig Vet Soc Cong 2004; p. 289.
22 Veenhuizen MF, et al. Proc 15th Int Pig Vet Soc Cong 1998; p. 264.
23 Chang WL, et al. Vet Rec. 1997;141:103.
24 Fellstrom C, et al. Proc 16th Int Pig Vet Soc Cong 2002; p. 192.
25 Guedes RMC. Vet Microbiol. 2003;91:135.
26 Guedes RMC. Vet Microbiol. 2003;93:159.
27 McCluskey T, et al. Infect Immun. 2002;70:2899.
28 Holyoake PK, Cutler RS. Aust Vet J. 1995;72:253.
29 Smith D, et al. Vet Rec. 1998;142:690.
30 Bronsvoort M, et al. J Swine Hlth Prod. 2001;9:285.
31 Just SD, et al. Swine Hlth Prod. 2001;9:57.
32 Bane, D. Proc 15th Int Pig Vet Soc 1998, p. 107.
33 Kroll J, et al. Proc 18th Int Pig Vet Soc Cong 2004; p. 149.
34 Guedes RMC, et al. Proc Am Ass Swine Vet. 2004:439.
35 McOrist S, et al. Res Vet Sci. 1995;59:255.
36 Lawson GHK, et al. Vet Microbiol. 1995;45:339.
37 Jasni S, et al. Vet Microbiol. 1994;41:1.
38 McOrist S, et al. Vet Rec. 1994;134:331.
39 Joens A.L., et al. Am J Vet Res. 1997;58:1125.
40 McOrist S, et al. Int J Syst Bacteriol. 1995;45:820.
41 McOrist S, et al. J Comp Pathol. 1996;115:35.
42 Quironi A, et al. Am J Physiol Cell Physiol. 2000;279:C1045.
43 McOrist S, et al. Infect Immun. 1993;61:4286.
44 MacIntyre N, et al. Vet Path. 2003;40:421.
45 Segales J, et al. Proc 16th Int Pig Vet Soc Cong 2000; p. 60.
46 Jensen TK, et al. Proc 18th Int Pig Vet Soc Con 2004; p. 326.
47 Bermudez V, et al. Proc 16th Int Pig Vet Soc Cong 2000; p. 34.
48 Holyoake PK, et al. J Vet Diag Invest. 1996;8:181.
49 Gebhart CJ, et al. Vet Pathol. 1994;31:462.
50 Moller K, et al. Vet Microbiol. 1998;62:59.
51 Knittel JP, et al. Am J Vet Res. 1998;59:722.
52 Jones GF. Am J Vet Res. 1993;54:1585.
53 McOrist S, et al. Vet Microbiol. 1994;41:205.
54 Boye M, et al. Vet Path. 1998;35:153.
55 Guedes RMC, et al. Vet Rec. 2003;153:432.
56 Jensen TK, et al. Vet Rec. 1999;145:613.
57 Guedes RMC, et al. Proc 18th Int Pig Vet Soc Cong 2004; p. 250.
58 Jensen TK, et al. Proc 17th Int Pig Vet Soc Cong 2002; p. 217.
59 Driemeier D, et al. Acta Histochim. 2002;104:285.
60 Jones GF, et al. J Clin Microbiol. 1993;31:2611.
61 Sun DK, et al. J Vet Sci. 2000;1:33.
62 Jordan DM, et al. J Vet Diag Invest. 1999;11:49.
63 Guedes RMC, et al. J Vet Diag Invest. 2002;14:528.
64 Guedes RMC, et al. Proc 17th Int Pig Vet Soc 2002; p. 220.
65 Pejsak Z, et al. Med Wet. 2001;57:723.
66 Lindecrona RH, et al. J Clin Microbiol. 2002;40:984.
67 Beckler DC, et al. Proc Am Ass Swine Vet. 2003:81.
68 La T, et al. Proc 18th Int Pig Vet Soc Cong 2004; p. 283.
69 Knittel JP, et al. Am J Vet Res. 1998;59:722.
70 Guedes RMC, et al. Can J Vet Res. 2002;66:99.
71 Guedes RMC, et al. J Vet Diag Inv. 2002;14:400.
72 Boyce M, et al. Vet Path. 1998;3:153.
73 Jensen TK, et al. Eur J Vet Path. 1997;3:115.
74 McOrist S, et al. Vet Path. 1989;26:260.
75 Huerta B, et al. J Comp Path. 2003;129:179.
76 Guedes RMC, et al. Proc Am Ass Vet Lab Diag. 2001:117.
77 Stege H, et al. Prev Vet Med. 2001;50:153.
78 Johansen M, et al. Proc 17th Int Pig Vet Soc Cong 2002; p. 222.
79 McOrist S, et al. Vet Rec. 1999;144:202.
80 Veenhuizen MF, et al. Swine Hlth Prod. 1998;6:67.
81 McOrist S, et al. Am J Vet Res. 1997;58:136.
82 McOrist S, Morgan J Proc 15th Int Pig Vet Soc Cong 1998;3:111.
83 Winkelman NL. Proc 15th Int Pig Vet Soc Cong 1998;3:112.
84 Paradis MA, et al. Proc 18th Int Pig Vet Soc Cong 2004; p. 835.
85 Marsteller T, et al. Vet Ther. 2001;2:51.
86 Paradis MA, et al. Swine Hlth Prod. 2004;12:176.
87 Winne de R, Neirynck W. Proc 16th Int Pig Vet Soc Cong 2002; p. 207.
88 McOrist S, et al. Vet Rec. 2000;146:61.
89 Winkelman NL, et al. J Swine Hlth Prod. 2002;10:107.
90 Winkelman NL, et al. Proc Am Ass Swine Vet. 2002;33:139.
91 Tasker JB, et al. Proc 18th Int Pig Vet Soc Cong 2004; p. 256.
92 Ripley P. Proc 15th Int Pig Vet Soc Cong 1998; p. 115.
93 McOrist S, et al. Proc 15th Int Pig Vet Soc Cong 1998; p. 114.
94 McOrist S, et al. Vet Rec. 1996;139:615.
95 Winkelman NL, et al. Proc 16th Int Pig Vet Soc Cong 2000; p.197.
96 Waters D, et al. J Swine Hlth Prod. 2001;9:109.
97 Kyriakis SC, et al. Vet Rec. 1996;138:489.
98 Sumano LH, et al. Pig J. 1998;42:24.
99 Winkelman NL, et al. Proc Am Ass Swine Vet. 2003:136.
100 Kinsley K, et al. Proc Am Ass Swine Vet. 2000:275.
The disease is associated with Lawsonia intracellularis, an obligate intracellular Gram-negative bacterium associated with proliferative enteropathy in pigs, horses, hamsters, dogs, deer, rabbits, rats, and ratites.1 There is close similarity in DNA among isolates from a variety of species.2 Isolates derived from pigs are infective in hamsters but there is no information regarding infectivity of porcine strains for horses, or vice versa.1 The organism from pigs is viable and able to produce disease in pigs when stored in feces at room temperature for up to 2 weeks.14
The disease was initially reported from North America and has recently been diagnosed in Australia and Europe.4,5,15 Nine of 164 randomly selected weanlings in the Hunter Valley of Australia had serum antibodies to L. intracellularis.5
The disease in foals occurs as isolated cases and as outbreaks on breeding farms.4 There is evidence that outbreaks begin after introduction of foals or weanlings to farms with no history of the disease, although whether this is coincidence or represents the mechanism of introduction of infection to the farm is unknown.4 Morbidity among foals and weanlings on affected farms is 20–25%, although this is based on disease outbreaks on only two farms.4 Case fatality rate is 15–20%.4
Affected foals are usually 3–13 months of age and disease in adults is rare. There is insufficient information to determine if there is a breed predisposition to the disease. The disease is presumably transmitted by the fecal–oral route, as in pigs, but there are no reports of experimental induction of the disease in foals.
The pathogenesis of the disease in foals has not been determined, but it is probably similar to that of the disease in pigs.6 Infection results in development of an enteropathy characterized by proliferation of intestinal crypt epithelial cells and infiltration of the lamina propria with mononuclear inflammatory cells.4,7-9 Subsequent malabsorption of small intestinal contents and protein loss from diseased intestine cause weight loss and hypoproteinemia characteristic of the disease in foals. However, there was no evidence of decreased absorption of glucose in three foals in which this was examined.4 Colic and diarrhea result from intestinal dysfunction and malabsorption. Hypoproteinemia and the subsequent decrease in plasma oncotic pressure result in edema and signs of hypovolemia. Death is associated with severe hypoproteinemia, inanition, and colic.
The disease may present as one with a short course characterized by rapid weight loss, colic and death within 2–3 days of onset of clinical signs or as a more chronic disease characterized by gradual development of weight loss and depression.4,7,8,10 Weight loss and poor body condition are consistent findings among foals affected by the chronic disease. Most affected foals have diarrhea that ranges in severity from acute profuse watery diarrhea to, more commonly, excessively soft feces. Foals are often depressed although they continue to nurse. Edema of the ventral abdomen and intermandibular space is common. Fever is not a consistent feature of the disease.
Ultrasonographic examination of the abdomen reveals multiple loops of mildly distended small intestine with thickened walls. Loops of intestine can have walls of 5–8 mm thick (normal <3 mm).
Many affected foals, and especially those that die of the disease, have concurrent diseases including parasitism and pneumonia.
The incubation period in pigs is 2–3 weeks but that in horses is unknown. Foals that recover from the disease may take several weeks to regain normal body weight. There do not appear to be long-term consequences of the disease in recovered foals.
Hypoproteinemia with moderate to severe hypoalbuminemia is present in most affected foals. Serum albumin concentrations can be as low as 0.6 g/dL (6 g/L).8 Hyperfibrinogenemia and mild anemia are common but not consistent findings.4 White cell count is elevated (>14 × 109 cells/L) in most foals. Serum sodium and chloride concentrations are lower than normal and serum creatinine concentrations higher than normal in about 50% of affected foals. PCR examination of feces for L. intracellularis is specific for detection of organism in affected foals. An indirect immunofluorescent assay detects serum IgG antibody to L. intracellularis in foals, although the specificity of this finding for detection of the disease in foals is unknown.4,5 Foals with proliferative enteropathy have titers of 1:30 or greater.5 Antemortem diagnosis in pigs is based on positive serology and detection of L. intracellularis DNA in feces by PCR.11
Gross lesions are mainly thickening and irregular corrugation of the small intestine.4,7,8 There is proliferation of intestinal crypt epithelium with projection of crypt cells into the intestinal lumen. The lamina propria is infiltrated by mononuclear inflammatory cells. Silver staining of intestinal sections reveals numerous short, curved bacteria in apical cytoplasma of crypt epithelial cells.4
Antemortem diagnosis of proliferative enteropathy in foals should be based on the presence of characteristic clinical, hematological and biochemical signs, positive serology and detection of L. intracellularis DNA in feces by PCR.
The primary differential diagnosis is parasitism by Parascaris equorum, Cyathostomes, and large strongyles (in older foals). Examination of feces for helminth ova is diagnostic in cases with patent infections, but parasite infestations are often not patent in young foals. A history of an adequate parasite control program makes parasitism less likely, but does not rule it out. Malnutrition due to inappropriate or inadequate feeding practices or agalactia should be ruled out as a cause of failure to thrive.
Protein-losing enteropathy secondary to enteritis and colitis may be associated with Salmonella sp., Rhodococcus equi, or Cryptosporidium sp. Other intestinal diseases that cause enteritis but less commonly cause protein loss include the intestinal clostridioses, equine granulocytic ehrlichiosis, and Bacteroides sp. infection. Intra-abdominal abscesses associated with R. equi or Streptococcus sp. can cause chronic weight loss and hematological signs similar to proliferative enteropathy.
Hypoproteinemia can occur secondary to gastrointestinal ulceration. Neoplasia is rare in foals of this age but intestinal lymphosarcoma can cause hypoproteinemia and weight loss. Intoxication by non-steroidal anti-inflammatory drugs can cause a protein-losing enteropathy.
Diarrhea and ill thrift caused by colitis and typhlitis associated with Brachyspira sp. (a spirochete) is reported from Japan.13
Principles of treatment are eradication of infection and correction of hypoproteinemia. Administration of antibiotics is curative in many foals.4,16 Isolates of the organism from pigs are sensitive in vitro to a wide range of antimicrobials including penicillin, erythromycin, difloxacin, virginiamycin, and chlortetracycline.14 Antibiotics used to treat L. intracellularis infection in foals include oxytetracycline (6.6 mg/kg q 12 h IV),16 doxycycline (10 mg/kg q 12 h, orally),16 chloramphenicol (50 mg/kg, q6 h, orally), or erythromycin estolate or similar product (15–25 mg/kg q 6–8 h orally), sometimes in combination with rifampin (5–10 mg/kg q 12 h orally). Erythromycin or oxytetracycline/doxycycline appear to be effective in treatment of affected foals.4,16 Chloramphenicol is used in place of erythromycin in foals that develop intractable or severe diarrhea when treated with erythromycin, but its use is illegal in some countries and is not recommended because of the risk of aplastic anemia in people exposed to the drug. Enrofloxacin might be effective, based on MIC values, but should be reserved as a drug of last resort because of the arthropathy associated with its use in foals.
Mildly or moderately affected foals require only administration of antimicrobials and nursing care. More severely affected foals may need intensive supportive care including intravenous administration of plasma and/or hetastarch to restore plasma oncotic pressure and minimize edema formation, fluid and electrolyte supplementation because of hypovolemia and abnormalities in serum electrolyte concentration, calorie enhanced diets or parenteral nutrition, and antiulcer medications if signs of gastric ulceration are present.
Specific control measures to prevent spread of the disease among horses have not been developed. Given the putative fecal–oral cycle of infection and association of outbreaks of the disease in pigs after introduction of new stock or mingling of groups, hygiene measures that minimize fecal contamination of the environment by potentially infected foals is sensible. The organism from pigs can survive in feces for up to 2 weeks. Foals with the disease should be isolated from healthy foals, although the duration of this isolation is not known, and should not be transported to other farms until clinical and hematological signs of the disease have resolved. The role for wildlife hosts, if any, in the disease of foals is unknown. There is currently no vaccine for the disease in foals.
1 Lawson GHK, Gebhart CJ. J Comp Path. 2000;122:77-100.
2 Cooper DM, et al. Vet Microbiol. 1997;54:47-62.
3 Collins AM, et al. Swine Health Prod. 2000;8:211-215.
4 Lavoie JP, et al. Equine Vet J. 2000;32:418-425.
5 McClintock SA, Collins AM. Aust Vet J. 2004;82:750-752.
6 Smith DGE, Lawson GHK. Vet Micro. 2001;82:331-345.
7 Frank N, et al. Equine Vet J. 1998;30:549-552.
8 Brees DJ, et al. JAVMA. 1999;215:511-514.
9 Williams NM, et al. J Vet Diagn Invest 8:254–256.
10 Schumacher J, et al. J Vet Int Med. 2000;14:630-632.
11 Knittel JP, et al. Am J Vet Res. 1998;59:722-726.
12 Smith DGE. Equine Vet J. 1998;30:452-453.
13 Shibahara T, et al. J Vet Med Sci. 2002;64:633-636.
14 McOrist S, et al. J Clin Micro. 1995;33:1314-1317.