359Chapter 42 Systemic Clostridial Infections

42 

Systemic Clostridial InfectionsDebra C. Sellon and Simon F. Peek*

C H A P T E R 

Clostridial Myonecrosis

Simon F. Peek

Etiology

Clostridial myonecrosis in horses is most often caused by infec-tion with Clostridium perfringens,1,2 although sporadic cases have been described in association with other Clostridium spp., including C. septicum,1,3,4 C. chauvoei,1,2,5 C. novyi,1,6 C. ramosum, C. sporogenes,1 and C. fallax.7 The majority of cases in the litera-ture have been single-species infections, but mixed infections have been reported.1,3 These highly pathogenic clostridial organisms are gram-positive, spore-forming, anaerobic bacilli that can elaborate numerous potent exotoxins. Vegetative growth of these clostridial species is accompanied by produc-tion of dermonecrotizing and vasoactive toxins that lead to gas production, extensive tissue damage, and necrosis, as well as rapidly developing, life-threatening systemic toxemia.

Epidemiology

The prevalence of clostridial myonecrosis is low, and although the disease is sporadic, more cases appear to occur in certain regions. Many cases reported in the literature are from the northeastern1,2,4 and midwestern1,5,8 areas of the United States, but the disease may occur anywhere in the United States,3 Canada,9 Europe,10 and the Southern Hemisphere.7,11 Although species such as C. perfringens can frequently be cultured from the environment and soil wherever livestock are found, the means by which spores or vegetative organisms gain access to areas of affected soft tissue is not fully understood. Some of the species involved, including the most frequently isolated species, C. perfringens, can also be found within the gastrointestinal tract, in both the vegetative and the spore form, and some strains may be regarded as commensals. Spores of some clostridia (e.g., C. sporogenes, C. histolyticum) can be found in healthy equine muscle tissue, which suggests that after creation of appropriate

anaerobic conditions, these spores may vegetate and begin exponential growth.12 However, spores of species typically iso-lated from clinical cases of equine myonecrosis have not, as yet, been identified dormant within muscle tissue.

Most cases of equine myonecrosis temporally occur soon after parenteral injection of pharmacologic or biologic agents (≤48 hours) in the affected area of the body.1,2,9 The condition is more common in the cervical musculature,1-4,8,9 but occa-sional cases involving the gluteal muscles1,8,9 and rarely the caudal thigh musculature1 have been reported. Cervical and throatlatch lesions are sometimes encountered secondary to inadvertent perivascular leakage of pharmacologics intended for intravenous (IV) administration.1 Traumatic wounds have rarely been associated with the condition.1,9 A wide array of pharma-cologic and biologic preparations have been incriminated as inciting causes of clostridial myonecrosis, including nonsteroidal antiinflammatories,1,2,4,8,9 antihistamines,1,2,9 multivitamins,1,2,9 antipyretics,1,7,13 dewormers,2,9,14 vaccines, diuretics,1 and syn-thetic prostaglandins.13 The most frequently reported pharma-cologic agent associated with the development of clostridial myonecrosis is flunixin meglumine, with the most cases occur-ring in the cervical region.1-11,13,14

Pathogenesis

Most information regarding the pathogenesis of clostridial myonecrosis comes from rodent models of infection. Because identical species of clostridia, specifically C. perfringens and C. septicum, are the most commonly recognized causes of human clostridial myonecrosis, it is reasonable to assume that etiologic-specific pathogenic factors are comparable. During vegetative growth of C. perfringens, the primary toxin implicated in the disease process is alpha (α) toxin, a dermonecrotic toxin with both phospholipase C and sphingomyelinase activity.15 The definitive role that α-toxin plays during myonecrosis has been well established using controlled vaccine protection studies in which mice immunized against the C-terminal domain of the α-toxin were protected against lethal intramus-cular (IM) doses of the organism, whereas unvaccinated controls were not.16 Deletional mutants of C. perfringens in which the

*The authors acknowledge and appreciate the original contributions of these authors, whose work has been incorporated into this chapter.

360 Section 3 Bacterial and Rickettsial Diseases

Speciation after anaerobic culture by genetic or fluorescent antibody techniques provides definitive confirmation and may guide prognosis.1,25

Real-time polymerase chain reaction (PCR) assays have been described for detection and differentiation of C. chauvoei and C. septicum.26,27 These rapid techniques may be used for ante-mortem diagnoses in large animals, which has largely been unrewarding in the past because bioassay techniques and direct florescent antibody techniques are restricted to postmortem samples. Early identification of anaerobic versus aerobic infec-tion may allow earlier intervention of appropriate antimicrobial therapy.

Therapy

Higher survival rates are associated with aggressive combina-tions of medical and surgical treatment.1,2,4 Barotherapy has become a component of the approach to treatment of human cases of clostridial myonecrosis, but hyperbaric oxygen cham-bers are only beginning to become available for use in large animal veterinary medicine. Rapid therapeutic intervention with high doses of IV crystalline penicillins should be consid-ered as soon as a presumptive diagnosis is made. Potassium penicillin at doses as high as 88,000 IU/kg every 2 hours (q2h) have been used, and conventional doses of 22,000 IU/kg q6h should be viewed as the minimum required dose, expense per-mitting.25 Oral metronidazole (25 mg/kg q6h) is often included in therapy but is unlikely to reach the high tissue levels that may be achieved with IV penicillin.

Intensive fluid, electrolyte, and cardiovascular support is indicated in the acute stages of clinical disease because dehydra-tion and systemic toxemia can be life threatening at this time.1-3,25 Many affected horses become hypotensive, with diminished cardiac output and renal function, complicated by substantial myoglobin release and the potential for pigment nephropathy. Fenestration of affected soft tissues appears to be an important part of therapy, and clinicians are advised to be aggressive in incising areas of subcutaneous emphysema, extending the incisions into deeper tissues and adjoining areas of healthy tissue (Fig. 42-2). Because of the obtunded menta-tion of many affected horses and the rapid progression of tissue necrosis in affected areas, these procedures can usually be per-formed under sedation without the need for local anesthesia.25 Frequent reevaluation of the horse for signs of spreading emphysematous cellulitis, necessitating repeated incisions and debridement, is advised. Local infusion of penicillin into tissue at the margins of debrided muscle may have some benefit in limiting the spread of the infection.

structural α-toxin gene has been removed are nonpathogenic, and virulence is restored by recombination with a plasmid expressing the wild-type gene.17 Other toxins that can be pro-duced during vegetative growth by C. perfringens and other clostridial species include theta toxin (perfringolysin), kappa toxin (collagenase), and mu toxin (hyaluronidase).15,18 Although other exotoxins elaborated by C. perfringens (and also other species) have highly potent and pathogenic effects extracellu-larly, no compelling evidence exists that they are required for lethal disease, as is the alpha toxin.

Many of the signs of systemic toxemia, cardiovascular col-lapse, and multiorgan dysfunction observed clinically in horses with clostridial myonecrosis can similarly be explained by observations made in rodent and rabbit models of gas gangrene. Alpha toxin directly suppresses myocardial contractility in vivo,19 whereas theta toxin is a potent reducer of systemic vascular resistance.20 Theta toxin has demonstrable ability to dysregulate polymorphonuclear/endothelial cell interactions, promoting leukostasis and interfering with normal cellular host responses to tissue injury at the active site of infection.21

Clinical Findings

Horses with clostridial myonecrosis demonstrate rapid soft tissue swelling, subcutaneous and deeper soft tissue emphy-sema, and rapid toxemia that may progress to circulatory collapse and multiorgan failure over just a few hours.1-3 Clini-copathologic data from horses that have died acutely corrobo-rate multiorgan failure and diffuse intravascular coagulation as two major pathologic processes that occur in terminally ill horses with clostridial myonecrosis.1,2,8 Some horses that develop clostridial myonecrosis have a recent history of colic, resulting in the administration of IM analgesics, with subse-quent myonecrosis in the region of the injection.1,2,8,9 Some authors have postulated that colic may truly be a prodromal sign of equine clostridial myonecrosis,4 drawing comparisons with the nausea and abdominal pain that are early symptoms of clostridial myonecrosis in human patients.22,23

Occasionally, horses with clostridial myonecrosis will develop hemolytic anemia/crisis after several days to 1 or 2 weeks of therapy.1,24 This appears to be a distinct entity to the life-ending, diffuse intravascular coagulation that other horses develop in the early stages of the disease. It is not certain whether hemo-lytic events are a direct effect of the clostridial infection. Some clostridia (e.g., C. septicum) can elaborate one or more exotoxins with in vivo hemolytic activity.22,23 Alternatively, hemolysis may be a potential immunologic complication of penicillin or other drug therapy.

Diagnosis

Clostridial myonecrosis may be presumptively diagnosed in any horse that develops acute-onset, rapidly progressive soft tissue swelling accompanied by emphysema in the area of a recent parenteral injection. Clostridial myonecrosis associated with penetrating traumatic wounds appears to represent a rare subset of cases.1,9 Acute-onset cellulitis without emphysema should be viewed as suspicious for the disease, but confirmed by cytologic evaluation and Gram stain before aggressive fasciotomy or debridement is performed (Fig. 42-1). Ultrasonography of areas of postinjection cellulitis should be performed to identify areas of deeper emphysema and gas production that may not yet be palpable in the immediate subcutis. Definitive etiologic confir-mation can be achieved by Gram stain and anaerobic culture of fluid aspirates from an affected soft tissue area. Numerous, characteristic gram-positive rods, sometimes with endospores present, can be easily visualized on air-dried smears, particularly from the periphery of an area of soft tissue emphysema.

Figure 42-1 Gram stain of aspirate from subcutaneous tissues of horse with acute clostridial myonecrosis. Notice the numerous, large, gram-positive rods.

361Chapter 42 Systemic Clostridial Infections

For horses that survive the acute stages of disease, prognosis will improve significantly. However, veterinarians should warn owners of the significant soft tissue and skin sloughing that will likely ensue over coming days to weeks (Fig. 42-3). Long-term wound care will often be needed, with many cases taking weeks to months before granulation and second-intention skin healing are complete. Cosmetically, some horses may heal with pigmen-tation changes and significant cicatrix formation (Fig. 42-4), but the visual appearance of healed wounds is often normal.25

Prognosis appears to vary according to the species of Clos-tridium involved. A much better prognosis is afforded to horses with soft tissue lesions associated with C. perfringens than to those with C. septicum or C. chauvoei infections.1 The largest retrospective study published to date reported an overall sur-vival rate of 73% for horses with clostridial necrosis when they were treated with a combination of aggressive medical and surgical therapy in a referral hospital setting. Horses with C. perfringens infection had a survival rate of 81%.1 There was no gender predilection observed in that study, but the disease did appear disproportionately to affect Quarter Horses. Previ-ous studies have demonstrated much higher case-fatality rates,2,8,9 and therefore clinicians and owners should be mindful of the need for prompt, aggressive, and potentially expensive therapy if horses are to have the best chance of survival. The majority of horses that die will succumb within the first few days.

Prevention

Although bacterin-toxoids should be in common use for pre-vention of disease caused by Clostridium tetani in all horses (see Chapter 44) and Clostridium botulinum in susceptible and at-risk foal populations (see Chapter 43), there is no standard vaccination practice for the prevention of clostridial myone-crosis in horses. Current preventive methods focus on appro-priate injection technique, particularly with respect to the location of IM administration of pharmacologic and biologic preparations. No protective effect appears to be gained from skin disinfection, hair clipping, or disinfection of the top of multidose vials before IM injection in preventing clostridial myonecrosis.26 When administering IM injections, however, particularly in the neck, where the disease most often occurs, it is prudent to ensure appropriate needle placement. When administering potentially irritant substances, particularly if lay people are responsible for injecting flunixin meglumine, it may be prudent to encourage use of the larger, better-vascularized, caudal thigh musculature.

Figure 42-2 Fasciotomy/myotomy incisions in gluteal region of 2-year-old Quarter Horse filly that developed clostridial myonecrosis secondary to vac-cination at the site. (Courtesy Dr. Susan Semrad.)

Figure 42-3 A, Mature Quarter Horse gelding showing skin and muscle sloughing 2 weeks after surgical fenestration of an area of clostridial myone-crosis in the cervical region. B, The same horse approximately 30 days after surgical fenestration showing near-complete granulation bed in prior area of clostridial myonecrosis.

A

B

Tyzzer’s Disease

Debra C. Sellon

In 1917, Ernest Edward Tyzzer28 described an infectious syn-drome of gastrointestinal and hepatic disease in Japanese Waltz-ing mice that came to be known as Tyzzer’s disease. He isolated and characterized the etiologic agent and reproduced the disease experimentally in mice. The etiologic agent was origi-nally named Bacillus piliformis. However, in 1994, Duncan et al29,30 demonstrated that the organism was more closely related to clostridial bacteria than to the Bacillus genus, and it was renamed Clostridium piliformis. In 1973, Swerczek et al31 described focal bacterial hepatitis in foals in Kentucky attribut-able to infection with Clostridium (Bacillus) piliformis.

Etiology

C. piliformis (also C. piliforme) is a motile, pleomorphic, gram-negative, spore-forming, obligate intracellular bacterium. It is classified as an organism that is “extremely oxygen sensitive” (EOS) and is gram positive only if fixation and staining are performed under strictly anaerobic conditions.31,32 C. piliformis

362 Section 3 Bacterial and Rickettsial Diseases

Serologic studies suggest that there is widespread exposure of horses to C. piliformis.98 Approximately 23% of horses tested had antibodies to the flagellar antigens of an equine C. piliformis isolate, 14% had antibody to epitopes of a rat isolate, and 5% had antibody to epitopes of a hamster isolate. This variability in seropositive rates between isolates suggests the possibility of multiple strains of bacteria that cause disease in horses.99,100 However, shared epitopes among strains have also been identi-fied,99 making it difficult to determine whether only certain strains may cause disease in animals or whether individual strains may be restricted to specific host species.

In a study of affected and unaffected Thoroughbred foals on a farm in California, several risk factors for Tyzzer’s disease were identified.37 Data from nine affected foals from a population of 322 foals were examined. In the final multivariable logistic regression analysis, foals born between March 13 and April 13 were 7.2 times as likely to develop Tyzzer’s disease as those

does not grow in cell-free media but can be propagated in the yolk sac of chick embryos or some types of cell culture. C. piliformis stains poorly with routine hematoxylin and eosin stains of formalin-fixed tissue samples. Silver impregnation or Giemsa stains facilitate visualization of the organism (Fig. 42-5). The large vegetative form of C. piliformis ranges from 8 to 40 mm in length. This vegetative phase is quite labile; in con-trast, spores may survive for up to 1 year in soiled bedding at room temperature or for 1 hour at 56° C (133° F).

Epidemiology and Pathogenesis

Infection with C. piliformis has been reported in a variety of lab-oratory, wild, and domesticated mammalian species, including the horse,30,33-49 cow,50,51 dog,52-55 cat,56 rat,57-63 mouse,58,59,64-69 hamster,69-71 gerbil,72-74 guinea pig,75-79 rabbit,80-82 muskrat,83-85 wombat,86 red panda,87 coyote,88 snow leopard,89 gray fox,90 raccoon,91 and serval.92 Among laboratory animals, clinical disease is most common in rabbits, gerbils, hamsters, and guinea pigs. Infections of mice and rats are more likely to be subclini-cal.62 There is one report of infection in a severely immuno-compromised person.93 There have been a few reports of infection in avian species.94,95 Disease has been reported in horses in many parts of the world, including North America, Australia, Europe, and Africa.

The natural route of infection of horses with C. piliformis is unknown, but the most likely route of exposure is by ingestion of spores from the environment. This theory is supported by the distribution of lesions in affected animals30,35,96 and experi-mental reproduction of disease in foals by oral transmission. Experimental infection of adult horses results in fecal shedding of the organism, and coprophagia may contribute to the likeli-hood of infection in foals.46 The sporadic nature of the disease in foals suggests that direct transmission is unlikely.50,97,98 However, clusters of disease may be observed on specific horse premises.30,37,46,47

Figure 42-5 Photomicrographs of the liver of foals with Tyzzer’s disease. Note the thin, filamentous bacilli visible within hepatocytes. A, Giemsa stain, 400×. B, Diff-Quik stain, 400×; imprint from liver of affected foal. C, Warthin-Starry (silver) stain, 160×. (Courtesy Dr. Charles Leathers.)

A

B

C

Figure 42-4 Scarring and skin depigmentation in a healed area of clostridial cellulitis and myonecrosis in throatlatch region associated with perivascular injection. (Courtesy Dr. Susan Semrad.)

363Chapter 42 Systemic Clostridial Infections

observed in parallel or random arrangements in hepatocytes at the periphery of these lesions if sections are silver-stained (Warthin-Starry stain). Occasionally, lesions consistent with enterocolitis or myocardial infection may be observed.39,41,103

Therapy

The prognosis for foals with Tyzzer’s disease is poor. Three foals with presumptive C. piliformis infection did survive.37,43 These foals received appropriate supportive care and antimicrobial therapy rapidly after they developed clinical signs. Because of difficulties cultivating C. piliformis in vitro, there is relatively little information regarding antimicrobial susceptibility pat-terns for this organism. The only available data are based on in vitro studies using embryonated eggs in which penicillin, tetracycline, erythromycin, and streptomycin were considered effective. Treatment of laboratory animals with either sulfon-amides or corticosteroids can induce active Tyzzer’s disease in carrier animals.104 A 10-day-old foal that survived presump-tive Tyzzer’s disease was treated with sodium penicillin and trimethoprim-sulfadiazine.43 In addition, the foal received intensive IV fluid therapy with dextrose, sodium bicarbonate, and potassium chloride solutions. Seizure activity was con-trolled with IM xylazine. Nutritional needs were met with IV parenteral nutrition. Additional supportive therapy included IV dimethyl sulfoxide and antiulcer prophylaxis. Clinical improve-ment was observed within 24 hours of initiation of therapy. Antimicrobial and antiulcer therapy was continued for approx-imately 3 weeks.

Prevention

No vaccines are available for prevention of Tyzzer’s disease in foals. Because C. piliformis is most likely transmitted by a fecal-oral route, good farm hygiene may be beneficial for decreasing the likelihood of disease. Maintaining foals in well-grassed pad-docks has been proposed as a preventive measure to decrease exposure to contaminated soil.45 It is also recommended that all foals receive adequate passive transfer of immune globulins soon after birth. High-risk foals37 should be closely monitored for the earliest signs of disease and treated aggressively as soon as these are recognized. Spores are reported to be sensitive to disinfection with 0.3% sodium hypochlorite105; however, this disinfectant is readily inactivated in the presence of organic matter and may not be an effective disinfectant for use in barns.

Public Health Considerations

There is a single report of human infection with C. piliformis in a severely immunocompromised patient.93

The complete reference list is available online at www. expertconsult.com.

born at other times; foals of nonresident (visiting) mares were 3.4 times more likely to be infected than resident foals; and foals of mares less than 6 years of age were 2.9 times as likely to develop disease as foals born to older mares. Seasonal risk may reflect management, environmental, or climatic factors that influenced disease incidence. The increased risk observed for foals born to nonresident mares and foals born to younger mares suggests that colostrum may be important for passive immunity in foals.37 Hook et al99 have demonstrated colostral transfer of antibodies to C. piliformis.

Very little is known about the pathogenesis of infection with C. piliformis in horses or other animals. Murine susceptibility to Tyzzer’s disease varies with host strain, age, and immune status. Depletion of neutrophils or natural killer cells in experimentally infected mice increases the severity of disease, but macrophage depletion does not alter the course of disease.101 In rodents and lagomorphs, outbreaks of disease are characterized by fatal diar-rhea, with pathology predominantly observed in the gastroin-testinal tract.102

Clinical Findings

Tyzzer’s disease affects foals between 7 and 42 days of age.37,43,47,48,98 Clinical signs include severe depression, fever, icterus, diarrhea, dehydration, and seizures. Some foals are found dead with no recognizable premonitory clinical signs. Almost all affected foals die; the overall course of disease is usually less than 48 hours from onset of clinical signs until death. Clinicopathologic abnormalities frequently include hemoconcentration, metabolic acidosis, hypoglycemia, hyper-bilirubinemia, and increased hepatic enzyme activities.37,43 Ultrasonographic examination of the abdomen of affected foals may reveal hepatomegaly with an increased vascular pattern.43

Diagnosis

Tyzzer’s disease should be considered as a differential diagnosis for foals between 7 and 42 days of age with compatible clinical signs and laboratory evidence of hepatitis. The likelihood of this diagnosis is increased if previous cases of C. piliformis infection have been confirmed on the premises. Currently, no reliable antemortem laboratory tests are available to confirm a diagnosis of Tyzzer’s disease in foals. Diagnosis is confirmed on postmor-tem examination by observation of typical gross and histo-pathologic lesions and observation of intracellular bacteria at the periphery of lesions when liver sections are stained appro-priately (see Fig. 42-5).

Pathologic Findings

At postmortem examination, affected foals have a grossly enlarged liver with multifocal, light-colored areas in the liver capsule and parenchyma.97 These light-colored areas corre-spond with areas of severe, random, diffuse acute to subacute hepatic necrosis.45 Intracellular filamentous bacilli can be

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363.e2 Section 3 Bacterial and Rickettsial Diseases

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363.e3Chapter 42 Systemic Clostridial Infections