- Host Animals
- Hosts/Species Affected
- Systems Affected
- Distribution Table
- List of Symptoms/Signs
- Disease Course
- Impact: Economic
- Zoonoses and Food Safety
- Disease Treatment
- Prevention and Control
- Links to Websites
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- swine dysentery
International Common Names
- English: swine dysentery, brachyspira hyodysenteriae, in pigs
Local Common Names
- USA: black scours; bloody scours
Pathogen/sTop of page Brachyspira hyodysenteriae
OverviewTop of page
Swine dysentery (SD) is a contagious diarrhoeic disease in pigs characterized by mucohaemorrhagic colitis and caused by strongly haemolytic spirochetes of the genus Brachyspira. The disease was originally described 80 years ago and is prevalent in most pig-rearing countries. For example, in the UK the disease was the second most commonly diagnosed enteric disease of pigs in 1995 (Taylor, 1995). Affected herds often suffer devastating production losses (Windsor and Simmons, 1981).
Swine dysentery was first described in 1921 in Indiana, USA by Whiting et al. (1921) as a bloody diarrhoea with necrotic haemorrhagic inflammation of the mucosa of the stomach and the large intestine. The disease has since been reported from most pig-rearing areas worldwide. The first observation in Europe dates from Italy in 1935 (Duthie, 1966). Swine dysentery was reported from Australia in 1938 (McLennan et al., 1938), The Netherlands in 1953 (Ulsen, 1953), Britain in 1957 (Birrel, 1957) and in Scandinavia in 1960 (Ronéus, 1960).
The disease appeared to increase in prevalence and severity with increasing intensification of pig industries and increasing trade in live pigs in the late 1960s. This stimulated the scientific community to investigate the then nebulous causes of SD. The true etiological factor was determined when Taylor and Alexander (1971) were able to provoke swine dysentery in pigs inoculated with a large spirochete.
A similar study was simultaneously performed by Harris et al. (1972a), who induced signs and lesions of dysentery in both specific pathogen-free (SPF) pigs inoculated with a large spirochete in combination with vibrio-like organisms, and in SPF pigs inoculated solely with the spirochete. The spirochete was named Treponema (T.) hyodysenteriae. Strain B78 was selected as the type strain of the new species.
Twenty years later, experiments capitalizing on advances in molecular biology, including DNA–DNA reassociation, sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS–PAGE) profiles of whole-cell proteins and analysis of 16S rRNA sequence data from strains of T. hyodysenteriae, all indicated that the organism was only distantly related to T. pallidum, the reference species of the genus Treponema (Stanton et al., 1991). Consequently, the genus name was changed, first to Serpula (Stanton et al., 1991) and then to Serpulina (Stanton, 1992).
The current designation of the classical etiological agent of SD, Brachyspira hyodysenteriae (Ochiai et al., 1997), was established when DNA-DNA reassociation experiments showed that the genuses Brachyspira and Serpulina should be designated to one genus, Brachyspira. According to taxonomic rules, the genus name Brachyspira (Hovind-Hougen et al., 1982) was selected because it had been proposed 11 years prior to Serpulina (Stanton, 1992). Recently, other Brachyspira spp. not identified as B. hyodysenteriae by laboratory methods have been associated with mucohemorrhagic diarrhoea in pigs including the proposed novel species “Brachyspira suanatina” (Råsbäck et al., 2007) and “Brachyspira hampsonii” (Chander et al., 2012). Experimental inoculation studies have confirmed the pathogenicity of these spirochetes through the induction of clinical disease and/or lesions indistinguishable from those associated with B. hyodysenteriae infection in mice (Burrough et al., 2012a) and susceptible pigs (Råsbäck et al., 2007; Burrough et al., 2012b; Rubin et al., 2013a). Accordingly, it has been recently concluded that the definition of SD be extended to include mucohemorrhagic colitis caused by any strongly haemolytic Brachyspira spp. (Hampson, 2014).
Taxonomy of Brachyspira hyodysenteriae
Brachyspira hyodysenteriae belongs to the genus Brachyspira (Fellström, 1996). Five officially recognized porcine species of Brachyspira have been described: B. hyodysenteriae; B. innocens (Kinyon and Harris, 1979); B.pilosicoli (Trott et al., 1996); Brachyspira intermedia (Stanton et al., 1997); and B. murdochii (Stanton et al., 1997). The genus Brachyspira is included in the order Spirochaetales, which is divided into three families Spirocaetaceae, Brachyspiraceae and Leptospiraceae. The family Spirocaetaceae contains the genera Borrelia, Brevinema, Cristispira, Spirochaeta, Spironema and Treponema, the family Brachyspiraceae contains the genus Brachyspira (Serpulina), and the family Leptospiracea contains the genera, Leptonema and Leptospira (Paster and Dewhirst, 2000). These organisms have morphological features in common which distinguish them from other bacteria. Spirochetes are unicellular, flexible and helical with one or more complete turns. Transmission electron microscopy has revealed that spirochetes have a coiled or wavy protoplasmatic cylinder, an outer sheath or tunica that envelops the whole organism, and unique organelles, called axial filaments (flagellae), which are inserted at each end of the cell and lie between the tunica and the cytoplasmatic membrane (Turner, 1976).
Host AnimalsTop of page
Hosts/Species AffectedTop of page
All domesticated pigs are probably susceptible to infection with B. hyodysenteriae and no breed disposition has been reported. Poor hygiene, cold temperatures, overcrowding, transportation and mixing of new stock may be predisposing factors (Griffin and Hutchings, 1980; Harris, 1984). Diet and immunological factors may also influence the clinical expression of SD. Clinical disease is mainly seen in growing, finishing or young adult stock.
Wild rodents are recognized as potential carriers of Brachyspira spp. (Joens and Kinyon, 1982) and are an important potential source of inter-farm and intra-farm spread. Numerous Brachyspira spp., including B. hyodysenteriae, have been recovered from rodents on pig farms (Backhans et al., 2010). The potential importance of waterfowl as a source of spirochete transmission between farms is supported by the recent isolation of “B. suanatina” in the faeces of mallard ducks in Sweden (Råsbäck et al., 2007) and “B. hampsonii” isolation from the faeces of migratory birds including lesser snow geese in Canada (Rubin et al., 2013b) and mallard ducks and graylag geese in Spain (Martínez-Lobo et al., 2013).
Systems AffectedTop of page digestive diseases of pigs
DistributionTop of page
Swine dysentery is a common disease worldwide (Roncalli and Leaning, 1976; Hampson et al., 1997; Hampson, 2000). However, very limited accurate information is available regarding the current prevalence of SD in different countries. Studies performed in the UK and Denmark have suggested a general SD prevalence of around 10% of herds (Møller et al., 1998; Thomson et al., 1998; Pearce, 1999) based on culture. A serological survey in the US indicated that 11% of swine herds had been exposed to B. hyodysenteriae (Mapother, 1993). An overall decline in the prevalence of the disease in the US probably occurred during the 1990s, due to the establishment of high health status herds and changes in management practices. Since the latter part of 2006, a re-emergence of SD has occurred in the United States and Canada (Burrough, 2013); however, the precise factors underlying this re-emergence are poorly understood and may include changes in feeding practices and other factors. The disease is likely to be present in most major pig producing countries outside Europe, the US and Australia, but is probably being controlled by the use of antimicrobial drugs (Hampson, 2000).
Distribution TableTop of page
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.Last updated: 10 Jan 2020
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Burundi||Absent, No presence record(s)||OIE Handistatus (2005)|
|Cabo Verde||Absent, No presence record(s)||OIE Handistatus (2005)|
|Central African Republic||Absent, No presence record(s)||OIE Handistatus (2005)|
|Congo, Democratic Republic of the||Absent, No presence record(s)||OIE Handistatus (2005)|
|Côte d'Ivoire||Absent, No presence record(s)||OIE Handistatus (2005)|
|Djibouti||Absent, No presence record(s)||OIE Handistatus (2005)|
|Egypt||Absent, No presence record(s)||OIE Handistatus (2005)|
|Eritrea||Absent, No presence record(s)||OIE Handistatus (2005)|
|Eswatini||Absent, No presence record(s)||OIE Handistatus (2005)|
|Ghana||Absent, No presence record(s)||OIE Handistatus (2005)|
|Guinea||Absent, No presence record(s)||OIE Handistatus (2005)|
|Libya||Absent, No presence record(s)||OIE Handistatus (2005)|
|Madagascar||Absent, No presence record(s)||OIE Handistatus (2005)|
|Mauritius||Absent, No presence record(s)||OIE Handistatus (2005)|
|Sudan||Absent, No presence record(s)||OIE Handistatus (2005)|
|Togo||Absent, No presence record(s)||OIE Handistatus (2005)|
|Tunisia||Absent, No presence record(s)||OIE Handistatus (2005)|
|Uganda||Absent, No presence record(s)||OIE Handistatus (2005)|
|Zimbabwe||Absent, No presence record(s)||OIE Handistatus (2005)|
|Bahrain||Absent, No presence record(s)||OIE Handistatus (2005)|
|Bhutan||Absent, No presence record(s)||OIE Handistatus (2005)|
|China||Present||CABI (Undated)||Present based on regional distribution.|
|-Fujian||Present||Guo JinHai et al. (1996)|
|Georgia||Absent, No presence record(s)||OIE Handistatus (2005)|
|India||Absent, No presence record(s)||OIE Handistatus (2005)|
|Indonesia||Absent, No presence record(s)||OIE Handistatus (2005)|
|Kazakhstan||Absent, No presence record(s)||OIE Handistatus (2005)|
|Kuwait||Absent, No presence record(s)||OIE Handistatus (2005)|
|-Peninsular Malaysia||Absent, No presence record(s)||OIE Handistatus (2005)|
|-Sabah||Absent, No presence record(s)||OIE Handistatus (2005)|
|Mongolia||Absent, No presence record(s)||OIE Handistatus (2005)|
|North Korea||Absent, No presence record(s)||OIE Handistatus (2005)|
|Oman||Absent, No presence record(s)||OIE Handistatus (2005)|
|Philippines||Absent, No presence record(s)||OIE Handistatus (2005)|
|Singapore||Absent, No presence record(s)||OIE Handistatus (2005)|
|Sri Lanka||Absent, No presence record(s)||OIE Handistatus (2005)|
|Syria||Absent, No presence record(s)||OIE Handistatus (2005)|
|Taiwan||Absent, No presence record(s)||OIE Handistatus (2005)|
|Thailand||Absent, No presence record(s)||Kramomtong et al. (1996); OIE Handistatus (2005)|
|United Arab Emirates||Absent, No presence record(s)||OIE Handistatus (2005)|
|Uzbekistan||Absent, No presence record(s)||OIE Handistatus (2005)|
|Belarus||Absent, No presence record(s)||OIE Handistatus (2005)|
|Croatia||Present||OIE Handistatus (2005)|
|Czechia||Absent, No presence record(s)||Cizek et al. (1996); OIE Handistatus (2005)|
|Estonia||Absent, No presence record(s)||OIE Handistatus (2005)|
|Finland||Present||Fossi (1996); OIE Handistatus (2005)|
|Germany||Present||1986||Waldmann et al. (2000)|
|Hungary||Present||Molnár (1996); OIE Handistatus (2005)|
|Liechtenstein||Absent, No presence record(s)||OIE Handistatus (2005)|
|Netherlands||Present||OIE Handistatus (2005)|
|North Macedonia||Absent, No presence record(s)||OIE Handistatus (2005)|
|Portugal||Absent, No presence record(s)||OIE Handistatus (2005)|
|Romania||Absent, No presence record(s)||OIE Handistatus (2005)|
|Slovakia||Absent, No presence record(s)||OIE Handistatus (2005)|
|Slovenia||Absent, No presence record(s)||OIE Handistatus (2005)|
|Spain||Absent, No presence record(s)||OIE Handistatus (2005)|
|Sweden||Present||Fellström et al. (1998); OIE Handistatus (2005)|
|United Kingdom||Present||Thomson et al. (1998); Pearce (1999); OIE Handistatus (2005)|
|-Northern Ireland||Present||OIE Handistatus (2005)|
|Barbados||Absent, No presence record(s)||OIE Handistatus (2005)|
|Bermuda||Absent, No presence record(s)||OIE Handistatus (2005)|
|British Virgin Islands||Absent, No presence record(s)||OIE Handistatus (2005)|
|Canada||Present||OIE Handistatus (2005)|
|-Quebec||Present||Harel et al. (1994)|
|Cayman Islands||Absent, No presence record(s)||OIE Handistatus (2005)|
|Cuba||Present||OIE Handistatus (2005)|
|Curaçao||Absent, No presence record(s)||OIE Handistatus (2005)|
|Dominica||Absent, No presence record(s)||OIE Handistatus (2005)|
|Guatemala||Absent, No presence record(s)||OIE Handistatus (2005)|
|Haiti||Absent, No presence record(s)||OIE Handistatus (2005)|
|Honduras||Absent, No presence record(s)||OIE Handistatus (2005)|
|Jamaica||Absent, No presence record(s)||OIE Handistatus (2005)|
|Mexico||Absent, No presence record(s)||OIE Handistatus (2005)|
|Nicaragua||Absent, No presence record(s)||OIE Handistatus (2005)|
|Panama||Absent, No presence record(s)||OIE Handistatus (2005)|
|Saint Kitts and Nevis||Absent, No presence record(s)||OIE Handistatus (2005)|
|Saint Vincent and the Grenadines||Absent, No presence record(s)||OIE Handistatus (2005)|
|United States||Present||Hampson et al. (1997); Hampson (2000); OIE Handistatus (2005)|
|-Iowa||Present, Widespread||Mapother (1993)|
|Australia||Present||Hampson et al. (1997); Hampson (2000); OIE Handistatus (2005)|
|-Western Australia||Present, Widespread||Mhoma et al. (1992)|
|French Polynesia||Absent, No presence record(s)||OIE Handistatus (2005)|
|New Zealand||Present||OIE Handistatus (2005)|
|Samoa||Absent, No presence record(s)||OIE Handistatus (2005)|
|Vanuatu||Absent, No presence record(s)||OIE Handistatus (2005)|
|Argentina||Present||OIE Handistatus (2005)|
|Bolivia||Absent, No presence record(s)||OIE Handistatus (2005)|
|Brazil||Absent, No presence record(s)||OIE Handistatus (2005)|
|-Rio Grande do Sul||Present||Barcellos et al. (2000)|
|Chile||Absent, No presence record(s)||OIE Handistatus (2005)|
|Colombia||Absent, No presence record(s)||OIE Handistatus (2005)|
|Ecuador||Absent, No presence record(s)||OIE Handistatus (2005)|
|Falkland Islands||Absent, No presence record(s)||OIE Handistatus (2005)|
|Guyana||Absent, No presence record(s)||OIE Handistatus (2005)|
|Uruguay||Absent, No presence record(s)||OIE Handistatus (2005)|
PathologyTop of page
Lesions are limited to the large intestine. Early lesions are first noted in the centrifugal and centripetal coils near the apex of the colon. Over time the changes may extend to the whole colon, and beginning day 4 may also affect the caecum. In some instances the whole large intestine may become involved. Typical changes in the acute stage of SD include hyperemia and oedema of the walls and mesentery of the large intestine. The mesenteric lymph nodes may be swollen, and colonic submucosal glands are often more prominent than normal. Mucus, and fibrin containing flecks of blood usually cover the mucosa. As the disease progresses, mucosal lesions may become more severe with increased fibrin exudation, and may form thick, mucofibrinous pseudomembranes. Once the lesions become chronic, the mucosa may take on the appearance of having marked superficial necrosis (Harris and Lysons, 1992).
Microscopically, the damage to the colonic mucosa and submucosa is characterized by superficial necrosis, eroded epithelium, oedema, leukocytic infiltration and hyperplasia of goblet cells. In acute SD, the mucus bilayer in the spiral colon is significantly altered with a reduction in sulphated mucins, decreased expression of mucin 4, and increased expression of mucin 5AC (Wilberts et al., 2014a). As the disease progresses, clumps of epithelial cells may detach from the lamina propria, resulting in exposure of capillaries followed by focal areas of haemorrhage. Large spirochetes are found in the lumen, within crypts and in the lamina propria and are readily visible by silver staining; however, this method is nonspecific and pathogenic spirochetes can be identified to the species level using in situ hybridization with oligonucleotide probes targeting B. hyodysenteriae (Boye et al., 1998) and “B. hampsonii” (Burrough et al., 2013). Using transmission electron microscopy (TEM), B. hyodysenteriae cells can be visualized invading epithelial cells, goblet cells and the lamina propria (Harris and Lysons, 1992).
DiagnosisTop of page
The first clinical evidence of disease is usually soft, yellow to grey faeces, but the most consistent sign of SD is a blood stained, mucoid diarrhoea. As the disease progresses, appearance of white mucofibrinous strands in the stools is almost pathognomonic. Other signs may include an arched back, suggesting abdominal pain, and accompanying anorexia and fever. Swine dysentery is a severe diarrhoeal disease with high mortality in typical cases that are not treated. However, severity may vary, and the use of antibacterials as growth promoters may suppress clinical signs. If the disease is not treated, prolonged diarrhoea may lead to severe dehydration, followed by death. A SD diagnosis may be further indicated by typical findings at necropsy (see Pathology).
Infection with Lawsonia intracellularis, Brachyspira pilosicoli, Trichuris suis or salmonellosis may induce clinical signs that resemble SD.
Bacterial culture and biochemical tests have traditionally been the most common methods used for detection of B. hyodysenteriae, and for differentiation from other species of intestinal porcine spirochetes. The organisms grow well on blood agar plates. Inclusion of antibiotics in the growth media, for example, a combination of spectinomycin, vancomycin and colistin, reduces the growth of other colonic microorganisms and allows the b-haemolytic colonies of B. hyodysenteriae to be identified (Fellström and Gunnarsson, 1995; Jensen, 1997). Porcine intestinal spirochetes can be classified into a scheme according to their biochemical properties (see Table; after Fellström et al., 1999). On a genetic basis, the groups roughly correspond to the hitherto officially recognized species of the genus Brachyspira isolated from pigs (Pettersson et al., 1996), i.e. B. hyodysenteriae (group I), Brachyspira intermedia (group II), B. murdochii (group IIIa), B. innocens (group IIIbc) and B. pilosicoli (group IV). In the biochemical classification, Brachyspira spp. associated with swine dysentery are characterized by strong b-haemolysis and belong to group I (see Table); however, the variability in indole reactivity between SD-associated spirochetes suggests modification of this table may be necessary (i.e. group Ia and Ib) should these putative novel species gain official recognition and require clinical differentiation.
Differentiation of porcine Brachyspira species by biochemical reactions
1Ind = Indole production
2Hipp = Hippurate hydrolysis
3a-gal = alpha-galactosidase activity
4ß-glu = beta-glucosidase activity
* = variable indole reactivity has been reported for B. hyodysenteriae; however, “B. suanatina” isolates are indole positive and “B. hampsonii” isolates are indole negative.
In addition to culture, other techniques for diagnosis of B. hyodysenteriae infections have been developed, including serology and polymerase chain reaction (PCR). Serological tests have shown a lack of sensitivity and/or specificity when used in the field, and are not suited for the reliable detection of individual carrier pigs (Hampson, 2000). Recent studies have suggested that screening ELISAs run on meat juices may be useful in detecting herds infected with B. hyodysenteriae (Song et al., 2012). A number of PCR systems with high specificity have been described (Elder et al., 1994; Leser et al., 1997; Atyeo and Hampson, 1998; Atyeo et al., 1998); however, these were often of limited practical use due to sensitivity issues when applied directly to faecal samples. A duplex assay specifically designed to detect B. hyodysenteriae and B. pilosicoli in pig faeces (La et al., 2003) was adopted by many diagnostic laboratories due to ease of use and rapid turnaround of results relative to selective culture; however, this and other species-based PCR assays may fail to detect novel and/or atypical isolates and should therefore be used with caution.
From a diagnostic standpoint, culture remains a highly sensitive method for detecting Brachyspira spp.; however, it is less specific than PCR and requires additional biochemical or molecular testing for speciation. In many veterinary diagnostic laboratories, cultured spirochetes can be speciated at the molecular level by a variety of PCR assays and/or nox gene sequencing as well as at the protein level using matrix-assisted laser desorption ionization time-of-flight mass spectrometry (Calderaro et al., 2013; Warneke et al., 2014).
Subspecies differentiation of Brachyspira spp. may be achieved by the use of multilocus enzyme electrophoresis (MEE; Lymbery et al., 1990), restriction endonuclease analysis (REA; Combs et al., 1989), restriction fragment length polymorphism (RFLP) analysis (Duhamel et al., 1992; Jensen and Stanton, 1993; Rohde et al., 2002), and pulsed field gel electrophoresis (PFGE; Atyeo and Hampson, 1998; Fellström et al., 1999).
A specific humoral immune response to outer membrane antigens of B. hyodysenteriae is induced following infection (Wannemuehler et al., 1988). Clinical disease is associated with the development of specific IgG, IgA and IgM antibodies in serum, and local production of IgA in gut mucosal tissues. Joens et al. (1984) suggested that the antibody secreted in the colon is the mechanism by which the recovered pig is protected against re-exposure to SD. Mucosal infection stimulates the production of memory cells, but humoral immunity alone is probably not responsible for the onset of a protective response to B. hyodysenteriae (Rees et al., 1989). Cell mediated immunity also may be important in protection against SD (Jenkins et al., 1982; Kennedy et al., 1992; Waters et al., 1999a). Repeated infection provides varying degrees of protection (Olson, 1974; Joens et al., 1979; Adachi et al., 1984; Rees et al., 1989).
List of Symptoms/SignsTop of page
|Digestive Signs / Anorexia, loss or decreased appetite, not nursing, off feed||Pigs:All Stages||Sign|
|Digestive Signs / Bloody stools, faeces, haematochezia||Pigs:All Stages||Diagnosis|
|Digestive Signs / Dark colour stools, faeces||Sign|
|Digestive Signs / Diarrhoea||Pigs:All Stages||Diagnosis|
|Digestive Signs / Melena or occult blood in faeces, stools||Sign|
|Digestive Signs / Mucous, mucoid stools, faeces||Pigs:All Stages||Diagnosis|
|General Signs / Dehydration||Pigs:All Stages|
|General Signs / Fever, pyrexia, hyperthermia||Pigs:All Stages||Sign|
|General Signs / Generalized weakness, paresis, paralysis||Sign|
|General Signs / Kyphosis, arched back||Pigs:All Stages||Sign|
|General Signs / Lack of growth or weight gain, retarded, stunted growth||Pigs:Piglet,Pigs:Weaner,Pigs:Growing-finishing pig,Pigs:Gilt||Sign|
|General Signs / Sudden death, found dead||Pigs:All Stages||Sign|
|General Signs / Underweight, poor condition, thin, emaciated, unthriftiness, ill thrift||Sign|
|General Signs / Weight loss||Pigs:All Stages||Sign|
|Pain / Discomfort Signs / Colic, abdominal pain||Pigs:All Stages||Sign|
Disease CourseTop of page
Many details of the pathogenesis of SD are still not understood. Infection occurs by ingestion of faecal material. The organism is protected from stomach acid by mucus in the dysenteric faeces (Taylor, 1995). Once ingested it invades the mucus and crypts of the mucosa in the large intestine and penetrates colonic enterocytes and goblet cells (Taylor and Blakemore, 1971). Early lesions can be observed before penetration, and therefore, penetration of cells is probably not a precondition for the initiation of SD (Wilcock and Olander, 1979; Albassam et al., 1985). Systemically, the disease is characterized by dehydration, acidosis and hyperkalemia, followed in severe cases by death.
Argenzio et al. (1980) demonstrated that the fluid losses in SD are exclusively the result of failure of the colon to reabsorb the endogenous secretions, due to a failure of the epithelial transport mechanisms to actively transport sodium and chloride from lumen to blood. Extensive fluid losses and ion imbalance occur with the disease, but B. hyodysenteriae does not invade the body nor produce a septicemic state (Kinyon et al., 1980).
Studies in gnotobiotic/germ-free pigs have indicated that B. hyodysenteriae is able to colonize the large intestine without the support of other microorganisms (Harris et al., 1972b; Meyer et al., 1974; Brandenburg et al., 1977; Whipp et al., 1982). However, several reports suggest that other colonic anaerobes act as supporting organisms (Harris et al., 1978; Whipp et al., 1982; Siba et al., 1994). Flagella and motility are probably involved in the colonization process (Kennedy and Yancey, 1996). The organism moves effectively at high speed by chemotaxis through viscous material such as mucin (Kennedy et al., 1988; Milner and Sellwood, 1994). Mutant strains of B. hyodysenteriae (Rosey et al., 1995; Kennedy et al., 1997), deprived of a flagellar gene, have been reported to be less virulent or avirulent in swine, which suggests that motility is an important virulence factor. Whether attachment is an important feature in the disease has not been conclusively demonstrated (Harris and Lysons, 1992).
It has been speculated (Savage, 1980) that an important virulence mechanism for pathogens of the intestinal tract could be their ability to utilize oxygen in order to colonize aerated environments such as the oxygen-respiring epithelial surfaces of the porcine large intestine. Stanton (1989) demonstrated that B. hyodysenteriae can metabolize substantial amounts of oxygen. Further mutants of B. hyodysenteriae strain B204 that have an inactivated NADH oxidase gene, and therefore a reduced ability to utilize oxygen, have been shown to be less virulent than the parent strain B204 (Stanton et al., 2000).
Interaction between B. hyodysenteriae and a fermentative bacterial flora in the colon and caecum has been suggested by Siba et al. (1994) and Pluske et al. (1996) to be involved in development of SD. Pigs fed a highly fermentative diet based on wheat and dehulled lupin seeds all developed SD in a challenge trial with B. hyodysenteriae, while pigs similarly challenged, but fed a diet based on boiled rice and animal protein were spared. The mean pH of caecal content in pigs before challenge for pigs fed the fermentative diet was 5.4, and 6.5 for pigs on the rice-based diet. Later studies (Durmic et al., 1998; Pluske et al., 1998) have further identified that high levels of non-starch polysaccharides, and starch which is not digested in the small intestine, are involved in the onset of SD. However, another similar study could not confirm the relationship between fermentable carbohydrates and the incidence of SD (Lindecrona et al., 2000).
Prohászka and Lukács (1984) demonstrated a pH-dependent antibacterial effect, attributed to the presence of volatile fatty acids (VFAs), in the contents of the large intestine of pigs. In vitro, a low pH <6.0) resulted in loss of motility of B. hyodysenteriae. They suggested that a diet based on maize silage is protective against SD because of the resulting acidic environment in the gut lumen of pigs.
Whether the synergism observed between B. hyodysenteriae and other anaerobes facilitates colonization or expression of pathogenicity (or both) has not been clearly demonstrated. Beckmann (1992) reported a CAMP-like phenomenon in Brachyspira isolates, when the isolates were streaked onto an isolate of Staphylococcus aureus. Whether such synergistic reactions could occur in vivo and if so, whether they might be of importance for pathogenicity is not known. An outer membrane Brachyspira protease may contribute to intestinal damage (Muniappa and Duhamel, 1997). Epithelial necrosis and vascular leakage late in the disease create conditions favouring overgrowth by opportunistic bacteria, which may be a contributing factor in pathogenesis (Albassam et al., 1985).
There is a close correlation between strong ß-haemolytic activity and pathogenicity among Brachyspira strains (Burrough et al., 2012b). However, the exact role played by the haemolysins is not known and the actual causes of the initial damage are not understood. The purified haemolysin (Kent et al., 1988) of B. hyodysenteriae has proved cytotoxic for a number of cell types, both in vitro and in vivo (Lysons et al., 1991; Hutto and Wannemuehler, 1999). Lysons et al. (1991) demonstrated damage to epithelial cells in ligated ileal and colonic loops of germ-free pigs injected with the haemolysin. The lesions, characterized by swelling and shedding of cells with disrupted organelles, were similar to those observed as early changes following injection of pig colonic loops with cultures of B. hyodysenteriae (Kang and Olander, 1990).
The importance of the haemolysin as a virulence factor has been emphasized by experimental infections in mice and pigs (ter Huurne et al., 1992a, b; Hyatt et al., 1994). These workers used recombinant DNA-techniques to produce mutants of B. hyodysenteriae deprived of the haemolysin tlyA gene. The mutants did not cause dysentery, but still produced mild lesions. From these experiments it seems reasonable to conclude that the haemolysin is an important factor in the pathogenesis of SD, though other factors may also be involved. However, reports of avirulent field isolates of B. hyodysenteriae have been published (Burrows and Lemcke, 1981; Lysons et al., 1982; Lee et al., 1993) and extended laboratory passages can produce strains that remain strongly b-haemolytic, but are no longer pathogenic for pigs (Hudson et al., 1974; Kinyon et al., 1977; Jensen and Stanton, 1993). On the other hand, potentiated pathogenicity by in vivo passages of B. hyodysenteriae in susceptible pigs has been demonstrated (Amtsberg et al., 1984; Blaha et al., 1984).
Other toxins may also be involved in causing lesions of SD (Baum and Joens, 1979; Nuessen et al., 1983; Greer and Wannemuehler, 1989a, b; Nibbelink et al., 1997) that may help provoke inflammatory lesions following infection with B. hyodysenteriae. However, Whipp et al. (1978) and Wilcock and Olander (1979) were unable to demonstrate toxic activity either with cell-free derivates of broth cultures or with sonically disrupted whole cultures of B. hyodysenteriae. Schmall et al. (1983) observed no increase above normal values of cyclic adenosine monosulphate (cAMP) and cyclic guanosine monophosphate (cGMP) in colonic mucosa of dysenteric pigs, suggesting that neither enterotoxin nor prostaglandins are involved in causing the diarrhoea.
EpidemiologyTop of page
Swine dysentery is caused by proliferation of strongly haemolytic Brachyspira spp. Infection occurs by ingestion of faecal material. The disease is usually introduced into a herd by carrier pigs, for example, animals that have recovered from a previous infection (Windsor and Simmons, 1981). Such pigs may continue to shed the infectious agent for months, without manifesting clinical signs (Songer et al., 1978; Fisher and Olander, 1981). In addition, B. hyodysenteriae has the ability to survive in pig faeces for up to 48 days (Chia and Taylor, 1978).
Although the infectious agent has only been demonstrated to occur as a natural infection in pigs and rheas (Jensen et al., 1996), wild rodents may be carriers of B. hyodysenteriae (Joens and Kinyon, 1982; Backhans et al., 2010) and waterfowl may also be a source of Brachyspira transmission between farms (Rubin et al., 2013b; Martínez-Lobo et al., 2013).
Brachyspira hyodysenteriae is recombinant with an epidemic population structure (Trott et al., 1997). As a consequence, it is unusal for more than one strain of B. hyodysenteriae to be present in a piggery (Fellström et al., 1999).
Impact: EconomicTop of page
Reduced production due to impaired growth, mortality, costs of medication and treatments, increased inputs for preventive measures and restrictions on movement of stocks may cause considerable economic losses. In Australia, losses resulting from reduced production and deaths was estimated to be 2.5% of annual gross national pig production (Hampson, 1991). Wood and Lysons (1988) estimated the cost due to decreased production in an infected herd in the USA to be US $12.60 per pig sold. In another infected herd in the USA, the costs of medication were estimated to be US $8.30 per pig marketed, and after eradication, medication costs were reduced to US $0.08 per pig marketed (Walter and Kinyon, 1990). Duhamel and Joens (1994) estimated the total national annual losses due to SD in the USA at US $115.2 million.
Zoonoses and Food SafetyTop of page
At present there is no evidence that swine dysentery occurs as a zoonotic disease. Consequently, it is unlikely that SD possesses any kind of threat to humans.
Disease TreatmentTop of page
A variety of antibacterials, such as tylosin, nitroimidazoles (dinitroimidazole, metronidazole, ronidazole), lincomycin, virginiamycin, tiamulin, olaquindox and carbadox, have been used successfully over the years in the treatment and prevention of SD. Although most of these drugs still may have some clinical effect, development of resistance for B. hyodysenteriae to several of those substances has been reported. Furthermore, the pharmacokinetic properties of olaquindox and carbadox make these antibacterials unsuitable for the treatment of SD (Graaf et al., 1988; Spierenburg et al., 1988), in spite of low minimum inhibitory concentrations (MICs) in vitro. Moreover, nitroimidazoles are no longer available in the EU and the USA. A number of reports indicate that pleuromutilins (tiamulin, valnemulin) are the only antimicrobials available with acceptable MICs in relation to pharmacokinetic properties for the treatment of Brachyspira (Rønne and Szancer, 1990; Binek et al., 1994; Buller and Hampson, 1994; Cizek et al., 1996; Fellström et al., 1996; Molnár, 1996; Karlsson and Franklin, 2000). Valnemulin sometimes produces adverse reactions (depression, ataxia, weakness, anorexia) when used in Danish and Swedish Landrace breeds and crosses.
Buller and Hampson (1994) claimed that having only one suitable substance for treatment of SD presents a potential threat to the pig industry. They suggested that the use of antimicrobials must be restricted to specific therapy. Accordingly, it is concerning that recent reports have shown tiamulin resistance in strains of B. hyodysenteriae from Spain (Hidalgo et al., 2011) and another report revealed a gradual rise in tiamulin MICs over time (Pringle et al., 2012). These resistant strains may be somewhat regional as a recent report of tiamulin susceptibility in Polish isolates showed no resistance (Zmudzki et al., 2012). Both the proposed novel Brachyspira spp. associated with SD appear similarly amenable to antimicrobial therapy as isolates of “B. suanatina” have been reported as susceptible to tiamulin, lincomycin, and valnemulin (Råsbäck et al., 2007) and North American isolates of “B. hampsonii” have been reported as sensitive to tiamulin, valnemulin, and carbadox (Mirajkar and Gebhart, 2013). Isolates of “B. hampsonii” from European waterfowl have also been reported as susceptible to tiamulin (Martínez-Lobo et al., 2013). Tiamulin water medication appears highly effective in resolving SD as the additional of tiamulin at 60 ppm in the water resolved clinical disease within 24 hours and eliminated spirochete shedding within 72 hours in pigs experimentally infected with either B. hyodysenteriae or “B. hampsonii” (Wilberts et al., 2014b).
The observation of resistant strains of B. hyodysenteriae underscores the importance of MIC testing of clinical isolates prior to treatment. Furthermore, these findings, and the limited number of efficacious antimicrobials available to treat SD, suggest that alternative means of controlling SD must be developed, such as all in-all out procedures, availability of SD-free stock, and eradication of strongly haemolytic Brachyspira spp. from affected herds.
Prevention and ControlTop of page
Immunization and Vaccines
Repeated infection provides protection in varying degrees. Vaccination has not been shown to offer any advantages over efficient treatment and control measures. Different approaches have been used in the development of vaccines. Most have involved the use of killed whole-cell preparations (Fernie et al., 1983; Diego et al., 1995; Waters et al., 1999b). Subunit vaccines and live vaccines have also been described (Wannemuehler et al., 1990; Lysons et al., 1986). All methods used have been shown to be only partly effective. A reverse vaccinology approach was used to identify B. hyodysenteriae proteins for use as recombinant vaccine components and revealed multiple potential immunogens that may prove useful (Song et al., 2009). No commercial vaccine against B. hyodysenteriae is currently available; however, autogenous bacterin vaccines have been used with some success.
Eradication of the infectious agent should always be considered in infected herds. If producers choose to 'live' with the disease rather than to try and eradicate it, all in-all out production combined with careful cleaning and disinfecting routines are the most important factors in the control of SD. Rodent control is also essential as wild rodents can serve as a source of spread within and between farms. To avoid introduction of SD and other diseases into herds, purchased animals should always be kept in quarantine for at least 3 weeks. If the SD status of such animals is unknown, treatment with pleuromutilins (tiamulin, valnemulin) should be performed.
ReferencesTop of page
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