GENERAL INTRODUCTION: SPIROCHAETES

Introduction

Swine dysentery (SD) in its typical acute disease manifestation is characterized by dysentery and the development of acute diphtheritic to necrotic enteritis with haemorrhage of the caecum and colon in weaned, growing, finishing and adult pigs. It results in temporary loss of condition, chronic wasting, or death. It is an important enteric disease in many countries.

The disease was first described in 1921 by Whiting and his co-workers in the USA who were able to reproduce it in transmission experiments using the gut contents of affected animals, thus proving that it was transmissible.⁴⁸ It was recorded in many countries of the world but the aetiological agent was unknown until 1971 when Taylor and Alexander described the isolation of a spirochaete and the reproduction of the disease by feeding pure cultures of it to pigs.⁴⁵ Shortly after that, Harris and his co-workers in the USA confirmed their findings and named the organism Treponema hyodysenteriae.^{14, 16} Both the organism and the disease have been extensively studied since then. This knowledge has been applied to methods used to diagnose, treat and control the disease, but has not yet resulted in a universally applicable vaccine.

Aetiology

The aetiological agents of SD are Brachyspira hyodysenteriae (formerly Serpulina or Treponema hyodysenteriae),³⁵ the more recently described  B. hampsonii⁷from North America, and B. suanatina in Northern Europe.³⁹ These species are included in a group of large oxygen-tolerant anaerobic spirochaetes found primarily in the large intestines of pigs and other mammals and birds. Brachyspira hyodysenteriae is 6 to l0 µm in length, flexible and active when viewed by phase contrast microscopy, and stains readily with aniline dyes such as carbol fuchsin. The cell contains a protoplasmic cylinder 350 nm in diameter with pointed ends and 7 to 14 fibrils inserting at each end and is surrounded by an envelope. Brachyspira hampsonii and B. suanatina are somewhat shorter in length and have 10-14 (B. hampsonii) and 7-8 (B. suanatina) flagella per cell end, respectively.^{33, 34}

The organisms can be grown on blood agar in atmospheres containing 5 per cent carbon dioxide and 95 per cent hydrogen. They form colonies 1 mm in diameter surrounded by β-haemolysis after 48 hours’ incubation.

Brachyspira hyodysenteriae is strongly β-haemolytic, indole positive, β-galactosidase negative, hippurate negative, and is antigenically distinct from other Brachyspira spp. With the exception of B. hampsonii being indole negative,³³ these features are shared with B. hampsonii and B. suanatina. It is sensitive to drying⁸ and acid conditions (pH <6,0), and is readily destroyed by heat but can survive in organic matter for days or weeks. Its specific antigens include 16 and 36 kDa proteins, 10 kDa envelope protein and a 46 kDa periplasmic flagellar protein.¹⁹

Historically, serotyping, multilocus enzyme electrophoresis (MLEE), 16S rRNA sequences, DNA:DNA hybridization, whole cell DNA probes, restriction enzyme analysis (REA), pulsed field gel electrophoresis, and recently, random amplification of polymorphic DNA (RAPD), matrix-assisted laser desorption ionization time-of-light spectrometry (MALDI-TOF), multi-locus sequence typing (MLST), multiple-locus variable-number tandem-repeat analysis (MLVA), and whole genome sequencing have all been used to identify/classify B. hyodysenteriae, but not B. hampsonii and B. suanatina.⁵¹

Epidemiology

Descriptions of the epidemiology and pathogenesis in this chapter are based on studies of B. hyodysenteriaeand not on B. hampsonii and B. suanatina. Swine dysentery occurs worldwide, is common in many European countries, and has recently been described as a re-emerging disease in for example the USA. However, official national incidence figures are scarce. Increasing numbers of herds are being founded using SD-free breeding stock housed in isolation. Dysentery-free pigs are being used in feeding enterprises, and separate site weaning and rearing operations are all reducing the prevalence of the disease in many countries.

Swine dysentery is transmitted to healthy, susceptible pigs by the ingestion of the faeces of affected or carrier animals or of material contaminated with their faeces.

Faecal shedding of the organism occurs during the incubation period of the disease (in the field this varies between 7 and 60 days but is usually 4 to 14 days in experimentally infected animals), its clinical course, and for up to 90 days following clinical recovery. Farms on which an outbreak of the disease has occurred remain infected unless depopulation and disinfection or a whole herd treatment have been carried out.⁴⁹  At least two electrophoretic types (ETs) or restriction endonuclease analysis (REA) types have been identified on individual farms, and B. hyodysenteriae appears to be a recombinant species with an epidemic population structure in which a few ETs have spread widely.⁴⁷

Pigs of all ages are affected although the peak prevalence is in weaned pigs of 6 to 12 weeks of age. The disease can be particularly severe in sows at farrowing or mid-lactation. It is usually introduced onto farms by the purchase of infected pigs, and spreads slowly from pen to pen by means of drainage channels or movement of infected pigs, by pig-to-pig contact, or by infected boots and implements.

The morbidity rate may reach 75 per cent of all pigs on the farm and mortality rates usually vary between 5 and 25 per cent. Some immunity results following recovery from infection with SD and recovered pigs rarely suffer relapses upon re-infection. Antibodies to B. hyodysenteriae have been identified in the sera, colonic secretions and colostrum of recovered animals.

Brachyspira hyodysenteriae can persist in moist faeces at 5 °C for up to 40 days and in faeces diluted 1:10 with tap water for up to 60 days. It can persist in slurry for at least six days and in soil for up to 41 days at −20 °C, 18 days at 14 °C, two days at 22 °C and less than five hours at 37 °C.⁸  Experimental transmission of the organism in infected soil to susceptible pigs is difficult to achieve. Drying or disinfectants such as formalin, phenols and hypochlorites at routine use dilutions eliminate the organism from contaminated pens within hours. Studies show that oxidizing disinfectants act within minutes.

Mice, rats, mallard ducks, crows, flies and dogs may be carriers of the organism in the field. Experimental infection of day-old chicks confirms that the domestic fowl could be involved in the epidemiology. Brachyspira hyodysenteriae isolated from mice on infected farms has been shown to be pathogenic for pigs, and the organism can be maintained in mouse populations. Brachyspira hampsonii and B. suanatina have been found in migratory waterfowl such as mallard ducks and geese,^{32, 39} and they may be involved in the spread of the infection.

Pathogenesis

Infection with B. hyodysenteriae is by ingestion. The organism passes through the stomach and small intestine, protected from stomach acid by mucus in dysenteric faeces in field cases, and reaches the large intestine.There, it invades the mucus and crypts of the mucosa within two hours of exposure. The organism colonizes the mucus, and moves in it by means of its flagellae. Inactivation of the flaA and flaB genes reduces motility.²⁴ It is attracted to substances such as sialic acid, L-fucose, L serine and blood in colonic mucus. Infection with B. hyodysenteriae alters the mucus layer with enhanced mucin production and disorganization of the mucus layer.³⁷ The bacteria multiply in the crypts and invade goblet and epithelial cells, damaging or disrupting them. Invasion through epithelial cell junctions may occur. Mature epithelial cells may be damaged by haemolysins and haemolysis-associated proteins, of which seven have been identified, among them tlyA, tlyB, tlyC and hlyA. It has been proven that tlyA negative mutants are of lowered pathogenicity compared to their wild types. The tlyB gene encodes a Clp protease, and tlyC encodes haemolysin C, both of which show haemolytic and cytotoxic activity in vitro. HlyA encodes for an acyl carrier protein (ACP) for acylation of toxins which may be needed for B. hyodysenteriae to display its haemolytic activity.⁵ However, many issues regarding the contribution of the haemolysins to the strong haemolysis caused by B. hyodysenteriae, B. hampsonii and B. suanatina are still unresolved.

An inflammatory response associated with mast cell degranulation and production of interleukin 6 develops. Inflammatory changes and superficial capillary dilatation and rupture in the large intestine result in loss of tissue and blood into the lumen as well as in failure of the colonic epithelium to re-absorb chloride and sodium ions,^{3, 42} which give rise to diarrhoea and faeces containing blood and mucus. Death is primarily due to dehydration, ion imbalances and toxaemia. A phagocytic response occurs within two to three days of the onset of clinical disease and specific serum antibodies can be demonstrated within seven to ten days of the onset of clinical signs and persist for up to 19 weeks. Serum antibodies may be related to protective immunity as pigs that have recovered from SD seem to be protected from reinfection.²⁰ IgG, IgA and IgM antibodies present in the lumen of the large intestine reduce the rate of mucosal colonization.⁴⁰ Furthermore, faecal IgA but not serum IgG was increased in pigs challenged with B. hyodysenteriae 17 days post infection.³¹ Glucose and lactate levels in serum are unaltered during the infection but gluconeogenic amino acids are decreased.²² Serum neutrophils that increase during infection could contribute to oedema and epithelial erosions.²¹ During recovery there is an increase in IL-10 and B. hyodysenteriae specific antibodies develop. Specific antigens include an outer-membrane associated lipoprotein SmpA⁴³ and a membrane lipoprotein BmpB.²⁷

Studies of B. hyodysenteriae infections in gnotobiotic pigs indicate that the organism on its own is capable of causing the disease. Lesions in conventional pigs are colonized by bacteria such as Campylobacter coli, Bacteroides vulgatus, Acetivibrio ethanolgignens, Fusobacterium necrophorum and other spirochaetes, which may aid in the development of the lesions. Only B. hyodysenteriae, however, will initiate the disease. The disease may be exacerbated by the presence of fermentable food residues reaching the caecum and colon, and, although disputed,³⁰ the severity of the clinical signs may be reduced experimentally by the use of totally digestible diets.⁴⁴  In addition, diets containing chicory root (includes inulin) help to prevent SD in experimentally challenged pigs.¹⁵

Clinical signs

The first signs of the disease in a group of pigs include restlessness, twitching of the tail, abdominal discomfort (demonstrated by the animal kicking its flanks), hollowing of the flanks, slight reddening of the skin and slight inappetence.¹ A transient fever of 40 to 40,6 °C may occur, but usually disappears at the onset of the diarrhoea, which persists throughout the course of the disease.

Diarrhoea may be the clinical sign first noted, particularly if affected animals are housed in pens fitted with solid floors, but the fluid faeces are much more difficult to detect in those with slatted floors or containing bedding comprising deep straw. Blood, mucus and, later, pieces of necrotic material appear in the faeces, which are yellowish at first but later become brownish-red, liquid in consistency and foul-smelling. When recovery begins after 7 to 14 days, the faeces revert to the normal colour and consistency but may contain large quantities of mucus.

Affected pigs show a rapid loss of bodily condition characterized by prominent ribs and backbones, a thin appearance, sunken eyes and a hollowness of the flanks (resulting from loss of colonic contents). The coat may be rough, the tail limp and the perineum stained with faeces, some of which may be mucoid or blood-stained. Affected pigs show a variable reduction in appetite but all continue to drink. The feed conversion rate and daily live weight gain may be permanently depressed. Permanently stunted pigs may pass khaki-coloured liquid faeces. In sows the reduced bodily condition may result in depression of reproductive performance.

Sucking piglets as young as three weeks of age may develop classic signs of the disease, although the peak prevalence is in weaned pigs of 6 to 12 weeks of age. The disease can be particularly severe in sows at farrowing or mid-lactation. Production depression on an affected farm may result in an extension of the rearing time from birth to 100 kg weight of up to 30 days as well as a 15 per cent increase in farm costs (as a result of the decrease in productivity) and treatment costs. The presence of passive or incomplete active immunity, a therapeutic and/or prophylactic treatment regimen of inadequate duration or level, and the use of antimicrobial growth promoters may all reduce the severity of the clinical signs and make them difficult to identify as being part of the SD syndrome. Other enteric diseases may occur concurrently and affect its clinical appearance or outcome.

Animals with chronic disease, low rectal temperatures (<38 °C) and chronic khaki-coloured diarrhoea should be killed on humane grounds.

Pathology

Pigs that have died of SD are usually in poor bodily condition, and the carcass may appear pale and faintly cyanotic. The changes typical of SD, which may be less specific in animals that have died of chronic disease, are best observed in recently affected pigs killed specifically for post-mortem examination.

On opening the peritoneal cavity it is frequently noticed that the mesenteric lymph nodes are pale and swollen, but are rarely congested. The large intestine is the only organ consistently affected, the lesion being, in acute cases, that of acute diphtheritic to necrotic typhlitis and colitis. It is usually flaccid and may be dark reddish-brown or congested and have a shiny surface due to oedema of the serosa.

Pale patches 1 to 2 mm in diameter may be seen under the serosal surface; these are hyperplastic lymphoid diverticula. The contents of the large intestine are fluid, foul-smelling and contain varying amounts of mucus, undigested food and necrotic material, and are brownish-red in colour. The appearance of the mucosa varies with the age of the lesion. In early cases it is swollen and congested, but subsequently becomes covered with blood-streaked mucus and patches of fibrin and diphtheritic material. There is an extensive layer of necrotic material in older lesions, which may become dislodged in patches to expose haemorrhagic mucosa.

In some cases the liver is swollen and friable and the gastric mucosa inflamed. The stomach contents may be fluid or contain fibrous material of dietary origin in straw-based systems.

Microscopic lesions in acute cases are confined to the large intestinal mucosa, which is inflamed. Much of the mucosal epithelium is shed and the underlying capillaries are congested. An exudate comprising cellular debris, fibrin, bacteria and red blood cells covers the exposed tissue and remaining epithelium, and oedema and an inflammatory cell infiltration into all parts of the intestinal wall are present. Spirochaetes can be visualized in the crypts of the colonic mucosa in the earliest lesions by microscopic examination of silver-stained sections, and these may be identified as B. hyodysenteriae by application of fluorescent antibody, peroxidase-linked antibody or by in situ hybridization using specific probes.

Diagnosis

Swine dysentery is diagnosed on the basis of the history of the outbreak indicating a slowly spreading enteric disease that follows the introduction of infected pigs, the clinical signs of blood and mucus in diarrhoeic faeces of affected pigs, and the post-mortem findings of mucohaemorrhagic to necrotic typhlocolitis. The presence of spirochaetes in Gram-stained air-dried or wet smears of faeces or of colonic lesions provides additional evidence.  Samples should be submitted as faecal swabs in transport medium, and colon contents or portions of colonic mucosa in sterile containers. In countries where postal services are efficient, they may be submitted by post.⁴⁶ Isolation of the organism may be attempted on selective media using spectinomycin blood agar containing 400 µg spectinomycin/ml or media containing additional antibiotics, but care should be taken in assessing the results as weakly haemolytic spirochaetes may also be isolated.²⁸ At present seven Brachyspira species are recognized in the colon of pigs, and faecal samples containing more than one species of Brachyspira are common. Therefore, care should be taken to obtain pure isolates. Haemolysis and isolation rates can be improved by slicing the agar or punching holes in it.³⁶

Diagnosis may then proceed by use of biochemical tests (see Table 1) and be confirmed by PCR-based assays. The exclusive use of PCR excludes minimal inhibitory concentration determinations.

Table 1 Diagnostic scheme of porcine Brachyspira spp. based on biochemical reactions (adapted from¹⁰)

PCR is usually performed on cultures, but can also be applied directly on faecal samples although with reduced sensitivity.³⁸ Most PCR systems are based on nox,⁴ 23S rDNA,²⁹ or the tlyA-genes,¹¹ and duplex,³⁸ and multiplex²⁶ PCRs have been developed with, for example, simultaneous detection of B. hyodysenteriae and B. pilosicoli. A drawback with the PCR based diagnostics is that they may fail to detect new genetic variants, e.g B. hampsonii² and B. suanatina.^{39, 41} Nox-PCR-based restriction fragment length polymorphism (nox-RFLP), is useful for the differentiation of all known pig Brachyspira species except B. hampsonii.⁴¹

Matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF) is used to some extent but failed to differentiate B. suanatina from B. hyodysenteriae.⁴¹

In conclusion, checking for strong haemolysis is the only reliable method of detecting Brachyspira strains causing clinical signs of SD, i.e. isolation remains necessary for a correct diagnosis, and is also required for susceptibility testing of isolates.

Serological tests such as microtitre agglutination and ELISA^{9, 25} can be used on a herd basis to identify infected herds.  Studies suggest that neither is 100 per cent sensitive or specific, and that neither can identify individual infected animals. The main problem is that these tests may detect antibody to common antigens present in non-B. hyodysenteriae spirochaetes and statistical interpretation may be required. Herd monitoring can be carried out using rectal faeces swabs or samples from recently weaned pigs for culture and/or PCR.

Differential diagnosis

The presence of fresh blood and mucus in the diarrhoeic faeces differentiates SD from most other enteric conditions. Salmonellosis is often associated with fever, the diarrhoeas of post-weaning colibacillosis, salmonellosis, and colitis due to other spirochaetes rarely contain thick mucus or frank blood, and transmissible gastroenteritis and epidemic diarrhoea caused by coronaviruses occur in explosive outbreaks rather than spreading slowly from pen to pen. Faecal blood in gastric ulceration and proliferative haemorrhagic enteropathy is altered and blackish. At post-mortem examination the restriction of lesions to the large intestine rules out all but diarrhoea due to other spirochaetes, which is a milder disease or infection. Trichuris suis and Oesophagostomum spp. infestations should be excluded.

Swine dysentery may occur with other diseases and then be atypical. In these situations, and following inadequate prophylactic or therapeutic drug treatment, laboratory examination is required for confirmation of the presence of B. hyodysenteriae.

Control

Treatment

In an outbreak individual severely affected animals should be treated by parenteral injection of a suitable therapeutic agent (see below). This is particularly important if feed medication is to be used. Medication of groups of affected animals is best done by administering drugs in the drinking water, but may be given in the feed. The antimicrobials given below may not all be available in any one country for regulatory reasons.

Pleuromutilins (tiamulin, valnemulin) are extremely effective in treatment of sensitive strains when given by injection (tiamulin) once a day for three days, in the drinking water (tiamulin) for five days , and in the feed (tiamulin, valnemulin) for 14 to 21 days. Resistance of B. hyodysenteriae to pleuromutilins is common in many countries, and Minimal Inhibitory Concentrations (MICs) of 4.0 mg/litre have been identified in isolates from a number of European countries and are associated with clinical resistance. Resistance to pleuromutilins is associated with several different, stepwise occurring mutations in the 23S rRNA gene. A new gene has been described, tva(A), encoding an ABC-F transporter, that does not lead to clinical resistance, but facilitates the development of higher-level resistance via mutations in ribosome genes.⁶ Pleuromutilins should not be given to pigs in conjunction with monensin, salinomycin or narasin as fatal ionophore poisoning may occur.

Lincomycin  may be given by injection once a day for three days, included in the drinking water and/or in the feed for 21 days.

Macrolides may be given by intramuscular injection twice a day (tylosin) for three days supplemented by medication in the drinking water (tylosin, tylvalosin) for seven to ten days, or in the feed (tylosin, tylvalosin) for 21 days.

Resistance to macrolides and lincomycin is widespread among B. hyodysenteriae isolates in most countries, but where organisms are sensitive, these antibiotics are effective. The genetic basis of macrolide and lincosamide resistance in B. hyodysenteriae is associated with a mutation in the nucleotide position 2058 of the 23S rRNA gene.²³ Resistance to macrolides develops very rapidly during treatment of B. hyodysenteriae.

All animals in drainage contact should be treated at therapeutic rates at the same time and their pens disinfected. They should, if possible, be moved to clean pens after treatment.

Prevention

The best way of prevention is to keep the herd “closed”. Extreme care should be taken regarding visitors, transport vehicles, rodents and other vectors, and there should be no purchase/recruitment of live animals.

The disease may also be prevented by administering a course of medication at therapeutic levels just after weaning and mixing, but should not be used as a routine measure due to the risk of development of antimicrobial resistance. Groups of treated pigs should then be housed in cleaned, dried and disinfected accommodation, and thereafter be maintained in isolation until slaughter. Maintenance in all-in-all-out systems is very effective and a rearing or finishing herd can be kept free from the disease even when the breeding stock are infected.

 No vaccines have as yet been registered for use worldwide. Killed whole cell vaccines made from one¹² or more serotypes and enriched with envelope proteins have been used in the field. They appear to reduce the prevalence of disease, to improve the performance of pigs, and to reduce the number of therapeutic treatments required to control it but do not prevent disease completely or prevent infection. Experimental vaccines composed of proteinase K digests of B. hyodysenteriae, subunits such as envelopes and flagellar proteins, live naturally attenuated organisms, and transposon mutants of the tlyA gene have all been shown to produce some degree of protection. A recent study¹⁷ describes marked serological heterogeneity of B. hyodysenteriae that has probably developed differently in diverse geographical areas over time. This situation represents a challenge for vaccine development since immunity to SD is serotype-specific.¹⁸ Autogenous vaccines are used in some countries but there is little evidence for their efficacy.

Management

General hygiene reduces the spread of the disease within a unit. The use of all-in-all-out husbandry, coupled with proper cleaning, disinfection and drying further helps to reduce the spread. Transmission from house to house can be reduced by disinfecting pathways and by using disinfectant foot dips and separate implements for each house. Slatted floors, solid partitions between pens, control of flies and rodents and lowering of slurry levels all reduce the spread of the disease. It is difficult, but not impossible, to control SD on earth floors or in earth runs. Drying and resting causes the organism to die out and disinfection of wooden pen furniture or walls and of earth floors can help free wooden pens or paddocks from residual organisms and allow restocking with clean pigs. Water for temperature control should be used carefully in piggeries as it is a common route for the transmission of the disease. Water used for sprays and pig pools should always come from a clean source and liquor from slurry systems should never be used to flush gutters or cool pigs. Pig pools should be for one pen of animals only and should drain directly into the slurry system. Pools should be drained and disinfected between batches.

Eradication

Eradication has been carried out in a number of ways.

Depopulation and restocking

The sale or slaughter of the infected stock is followed by cleaning and disinfection of pig houses, disposal of manure, an empty period to allow drying (it is best done in hot dry weather), and rodent and fly control. The clean unit is then restocked with SD-free pigs (usually originally hysterectomy-derived) from a breeding company and the herd is maintained in isolation except for the purchase of disease-free breeding stock or the purchase of semen. Such herds have a good chance of  remaining disease-free indefinitely, providing appropriate biosecurity measures are maintained regarding visitors, transport vehicles, and rodents/other vectors, Purchase/recruitment of live animals should be avoided even from officially declared SD free stock. Depopulation may take place as a single event, or the pigs may be moved to other holdings for finishing so as to reduce the period that the farmer is without income.

Treatment of the whole herd accompanied by disinfection and rodent control

A number of agents (tylosin, dimetridazole, ronidazole, lincocin and pleuromutilin) have been shown to eliminate B. hyodysenteriae.¹³ However, before treatment MIC-determinations of the SD causing agent should be performed. Treatment at the recommended rate (extended to double the usual period for safety) of the whole herd with an agent to which sensitivity has been established, followed by cleaning, disinfection and the isolation of all farrowing sows and their litters until both have undergone full periods of treatment, eliminates the disease.

Partial depopulation helps the cleaning process and allows for the increase in growth rate that follows successful eradication. Breeding stock should be purchased from disease-free herds.

Partial eradication accompanied by control

Where hygiene and management are good, the breeding stock may be treated and the organism eliminated. Treated pigs are isolated from the contaminated remnant of the herd by a disinfectant barrier along drainage lines which is moved forward ahead of the clean piglets being born.⁵⁰ Sufficient space for cleaning must be cleared by partial depopulation of the feeding herd and clinical disease should be suppressed by prompt treatment or continuous medication.

Such schemes are often cost effective, the cost being recouped within six months, and may reduce finishing periods by 7 to 21 days. Isolation and treatment of all incoming animals should be practised.

Isolation of the herd and the purchase of clean breeding stock is essential for continued success.

Monitoring for freedom from swine dysentery

A number of programmes have been devised to monitor the success of eradication campaigns as well as the continued freedom from infection status of pig-breeding companies. Herds to be monitored should not be subjected to the use of treatment regimens that can suppress SD, e.g. antimicrobial growth promoters should not be used.  Farm staff should monitor the herd for evidence of clinical signs of SD and report all outbreaks of diarrhoea for investigation, not merely those in which blood and mucus are present in the faeces of affected animals. Regular veterinary inspections that include inspection of slatted pens in which dysenteric faeces are difficult to identify and rectal faecal sampling followed by visual inspection confirms clinical freedom from infection. A regular programme of monitoring the post-mortem findings of any dead pigs helps to detect early or mild cases.

Ideally, this programme can be augmented by serology (if available) using blood samples from pigs slaughtered in an abattoir or those aged more than ten to 12 weeks. Inspection of colons after slaughter may be carried out. Bacteriological culture of material from colonic lesions from slaughtered pigs or from those that have died should be done on a regular basis. Diarrhoeic faeces or rectal faecal samples taken at regular inspections can be examined for strongly haemolytic Brachyspira spp. Animals with post-weaning diarrhoea, those recently mixed and young gilts at service are most likely to yield evidence of infection in a subclinically infected herd. Monitoring should be by culture and PCR, but the laboratory carrying out the examination must be capable of distinguishing between Brachyspira spp., as non-pathogenic species may not be eliminated by the treatments described.

References

1. ALEXANDER, T. J. L. & TAYLOR, D. J., 1969. The clinical signs, diagnosis and control of swine dysentery. The Veterinary Record, 85, 59–63.
2. ALLER-MORÁN, L. M., JAVIER MARTÍNEZ-LOBO, F., RUBIO, P. & CARVAJAL, A., 2016. Cross-reactions in specific Brachyspira spp. PCR assays caused by “Brachyspira hampsonii” isolates: implications for detection. Journal of Veterinary Diagnostic Investigation. 286, 755–759. doi: 10.1177/1040638716667528.
3. ARGENZIO, R. A., WHIPP, S. C. & GLOCK, R. D., 1980. Pathophysiology of swine dysentery: Colonic transport and permeability studies. Journal of Infectious Diseases, 142, 676–684.
4. ATYEO, R. F., STANTON, T. B., JENSEN, N. S., SURIYAARACHICHI, D. S. & HAMPSON, D. J., 1999. Differentiation of Serpulina species by NADH oxidase gene (nox) sequence comparisons and nox-based polymerase chain reaction tests. Veterinary Microbiology. 1, 47-60, doi:10.1016/S0378-1135(99)00030-9.
5. BELLGARD, M. I., WANCHANTHUEK, P., LA, T., RYAN, K., MOOLHUIJZEN, P., ALBERTYN, Z., SHABAN, B., MOTRO, Y., DUNN, D. S., SCHIBECI, D., HUNTER, A., BARRERO, R., PHILLIPS, N. D. & HAMPSON, D. J., 2009. Genome sequence of the pathogenic intestinal spirochete Brachyspira hyodysenteriae reveals adaptations to its lifestyle in the porcine large intestine. PloS One, 4(3), e4641.
6. CARD, R. M., STUBBERFIELD, E., ROGERS, J., NUNEZ-GARCIA, J., ELLIS, R. J., ABUOUN, M., STRUGNELL, B., TEALE, C., WILLIAMSON, S. & ANJUM, M. F., 2018. Identification of a new antimicrobial resistance gene provides fresh insights into pleuromutilin resistance in Brachyspira hyodysenteriae, aetiological agent of swine dysentery. Frontiers in Microbiology. 9, 1183. doi:10.3389/fmicb.2018.01183.
7. CHANDER, Y., PRIMUS, A., OLIVEIRA, S. & GEBHART, C. J., 2012. Phenotypic and molecular characterization of a novel strongly hemolytic Brachyspira species, provisionally designated Brachyspira hampsonii. Journal of Veterinary Diagnostic Investigation, 245, 903–910.
8. CHIA, S. P. & TAYLOR, D. J., 1978. Factors affecting the survival of Treponema hyodysenteriae in dysenteric pig faeces. Veterinary Record, 103, 68–70.
9. EGAN, I. T., HARRIS, D. L. & JOENS, L. A., 1983. Comparison of the microtitration agglutination test and the ELISA for the detection of herds affected with swine dysentery. American Journal of Veterinary Research, 44, 1323–1328.
10. FELLSTRÖM, C. & GUNNARSSON, A., 1995. Phenotypic characterisation of intestinal spirochaetes isolated from pigs. Research in Veterinary Science. 59, 1-4. doi: 10.1016/0034-5288(95)90021-7.
11. FELLSTRÖM, C., ZIMMERMAN, U. & ASPAN, A., 2001. The use of culture, pooled samples and PCR for identification of herds infected with Brachyspira hyodysenteriae. Animal Health Research Reviews, 21, 37–43.
12. FERNIE, D. S., RIPLEY, P. H. & WALKER, P. D., 1983. Swine dysentery: Protection against experimental challenge following single dose parenteral immunisation with inactivated Treponema hyodysenteriae. Research in Veterinary Science, 35, 217–221.
13. GLOCK, R. D., 1984. Eliminating swine dysentery from selected herds. Modern Veterinary Practice, 65, 611–614.
14. GLOCK, R. D. & HARRIS, D. L., 1972. Swine dysentery II. Characterisation of lesions in pigs inoculated with Treponema hyodysenteriae in pure and mixed culture. Veterinary Medicine, Small Animal Clinician, 67, 65-68.
15. HANSEN, C. F., PHILLIPS, N. D., LA, T., HERNANDEZ, A., MANSFIELD, J., KIM, J. C., MULLAN, B. P., HAMPSON, D. J. & PLUSKE, J. R., 2010. Diets containing inulin but not lupins help to prevent swine dysentery in experimentally challenged pigs. Journal of Animal Science. 10, 3327–3336. doi: 10.2527/jas.2009-2719.
16. HARRIS, D. L., GLOCK, R. D., CHRISTENSEN, C. R. & KINYON, J. M., 1972. Swine dysentery. I. Inoculation of pigs with Treponema hyodysenteriae (new species) and reproduction of the disease. Veterinary Medicine and Small Animal Clinician, 67, 61-68.
17. HERBST, W., SCHNEIDER, S., BALJER, G. & BARTH, S. A., 2017. An update of Brachyspira hyodysenteriae serotyping. Research in Veterinary Science. 111, 135-139. doi: 10.1016/j.rvsc.2017.02.015.
18. JOENS, L. A., HARRIS, D. L. & BAUM, D. H., 1979. Immunity to Swine dysentery in recovered pigs. American Journal of Veterinary Research, 40(10), 1352-1354.
19. JOENS, L. A. & MARQUEZ, R. B., 1986. Molecular characterisation of proteins from porcine spirochaetes. Infection and Immunity, 54, 893–896.
20. JOENS, L. A., NIBBELINK, S. & GLOCK, R. D., 1997. Induction of gross and microscopic lesions of porcine proliferative enteritis by Lawsonia intracellularis. American Journal of Veterinary Research, 58, 1125–1131.
21. JONASSON, R., ANDERSSON, M., RASBÄCK, T., JOHANNISSON, A. & J., J.-W. M., 2006. Immunological alterations during the clinical and recovery phases of experimental swine dysentery. Journal of Medical Microbiology, 55, 845-855. doi: 10.1099/jmm.0.46538-0.
22. JONASSON, R., ESSÉN-GUSTAVSSON, B. & JENSEN-WAERN, M., 2007. Blood concentrations of amino acids, glucose and lactate during experimental swine dysentery. Research in Veterinary Science, 3, 323-331. doi: 10.1016/j.rvsc.2006.08.005.
23. KARLSSON, M., FELLSTRÖM, C., HELDTANDER, M. U. K., JOHANSSON, K. E. & FRANKLIN, A., 1999. Genetic basis of macrolide and lincosamide resistance in Brachyspira (Serpulina) hyodysenteriae. FEMS Microbiology Letters, 2, 255–260. doi: 10.1111/j.1574-6968.1999.tb13476.x.
24. KENNEDY, M. J., ROSEY, E. L. & YANCEY, R. J., 1997. Characterization of flaA⁻ and flaB⁻ mutants of Serpulina hyodysenteriae: both flagellin subunits, FlaA and FlaB, are necessary for full motility and intestinal colonization. FEMS Microbiology Letters, 1, 119–128. doi: 10.1111/j.1574-6968.1997.tb10472.x.
25. KRAMOMTONG, I., NERAMITMANSOOK, W., WHIPP, S. C., JOENS, L. A. & LIMAWONGPRANEE, S., 1996. Comparison of ELISA and selective culture in the diagnosis of swine dysentery in Thailand. Veterinary Record, 138, 332–333.
26. LA, T., COLLINS, A., PHILLIPS, N., OKSA, A. & HAMPSON, D., 2006. Development of a multiplex PCR for rapid detection of the enteric pathogens Lawsonia intracellularis, Brachyspira hyodysenteriae, and Brachyspira pilosicoli in porcine faeces. Letters in Applied Microbiology, 42, 284-288. doi: 10.1111/j.1472-765X.2005.01831.x.
27. LEE, B. J., LA, T., MIKOSZA, A. S. J. & HAMPSON, D. J., 2000. Identification of the gene encoding BmpB, a 30 kDa outer envelope lipoprotein of Brachyspira (Serpulina) hyodysenteriae, and immunogenicity of recombinant BmpB in mice and pigs. Veterinary Microbiology, 3, 245-257. doi:10.1016/S0378-1135(00)00244-3.
28. LEMCKE, R. M. & BURROWS, M. R., 1981. A comparative study of spirochaetes from the porcine alimentary tract. Journal of Hygiene, 86, 173–182.
29. LESER, T. D., MOLLER, K. & JENSEN, T. K., 1997. Specific detection of Serpulina hyodysenteriae and potentially pathogenic weakly beta-haemolytic porcine intestinal spirochetes by polymerase chain reaction targeting 23S rDNA. Molecular and Cellular Probes, 115, 363–372.
30. LINDECRONA, R. H., JENSEN, T. K., JENSEN, B. B., LESER, T. D., JIUFENG, W. & MOLLER, K., 2003. The influence of diet on the development of swine dysentery upon experimental infection. Animal Science, 1, 82-87.
31. MAHU., 2017.
32. MARTÍNEZ-LOBO, F. J., HIDALGO, Á., GARCÍA, M., ARGÜELLO, H., NAHARRO, G., CARVAJAL, A. & RUBIO, P., 2013. First identification of "Brachyspira hampsonii" in wild European waterfowl. PLoS One, 8, 12, e82626. doi: 10.1371/journal.pone.0082626.
33. MIRAJKAR, N. S., PHILLIPS, N. D., LA, T., HAMPSON, D. J. & GEBHART, C. J., 2016. Characterization and recognition of Brachyspira hampsonii sp. nov., a novel intestinal spirochete that is pathogenic to pigs. Journal of Clinical Microbiology, 54, 2942–2949. doi: 10.1128/JCM.01717-16.
34. MUSHTAQ, M., ZUBAIR, S., RASBÄCK, T., BONGCAM-RUDLOFF, E. & JANSSON, D. S., 2015. Brachyspira suanatina sp. nov., an enteropathogenic intestinal spirochaete isolated from pigs and mallards: genomic and phenotypic characteristics. BMC Microbiology, 15, 208. doi: 10.1186/s12866-015-0537-y.
35. OCHIAI, S., ADACHI, Y. & MORI, K., 1997. Unification of the genera Serpulina and Brachyspira, and proposals of Brachyspira hyodysenteriae Comb. Nov., Brachyspira innocens Comb. Nov. and Brachyspira pilosicoli Comb. Nov. Microbiology Immunology, 41, 445–452.
36. OLSON, L. D., 1996. Enhanced isolation of Serpulina hyodysenteriae by using sliced agar media. Journal of Clinical Microbiology, 34, 2937–2941.
37. QUINTANA-HAYASHI, M. P., MAHU, M., DE PAUW, N., BOYEN, F., PASMANS, F., MARTEL, A., PREMARATNE, P., FERNANDEZ, H. R., TEYMOURNEJAD, O., VANDE MAELE, L., HAESEBROUCK, F. & LINDÉN, S. K., 2015. The levels of Brachyspira hyodysenteriae binding to porcine colonic mucins differ between individuals, and binding is increased to mucins from infected pigs with de novo MUC5AC synthesis. Infection and Immunity, 4, 1610-1619.
38. RASBÄCK, T., FELLSTRÖM, C., GUNNARSSON, A. & ASPÁN, A., 2006. Comparison of culture and biochemical tests with PCR for detection of Brachyspira hyodysenteriae and Brachyspira pilosicoli. Journal of Microbiological Methods, 662, 347-353.
39. RASBÄCK, T., JANSSON, D. S., JOHANSSON, K. & FELLSTRÖM, C., 2007. A novel enteropathogenic, strongly haemolytic spirochaete isolated from pig and mallard, provisionally designated ‘Brachyspira suanatina’ sp. nov.. Environmental Microbiology, 9, 983-991. doi: 10.1111/j.1462-2920.2006.01220.x.
40. REES, A. S., LYSONS, R. J., STOKES, C. R. & BOURNE, F. J., 1989. Antibody production in the pig colon during infection with Treponema hyodysenteriae. Research in Veterinary Science, 47, 263–269.
41. ROHDE, J., MAJZOUB-ALTWECK, M., FALKENAU, A., HERMANNS, W., BURROUGH, E. R., RITZMANN, M. & STADLER, J., 2018. Occurrence of dysentery-like diarrhoea associated with Brachyspira suanatina infection on a German fattening pig farm. Veterinary Record, 182, 195.
42. SCHMALL, L. M., ARGENZIO, R. A. & WHIPP, S. C., 1983. Pathophysiologic features of swine dysentery: Cyclonucleotide independent production of diarrhoea. American Journal of Veterinary Research, 44, 1309–1316.
43. SELLWOOD, R., WALTON, F., THOMAS, W., BURROWS, M. R. & CHESHAM, J., 1995. Expression of the SmpA outer membrane lipoprotein of Serpulina hyodysenteriae strain P18A in vivo. Veterinary Microbiology, 1, 25-35. doi: 10.1016/0378-1135(94)00108-9.
44. SIBA, P. M., PETHICK, D. W. & HAMPSON, D. J., 1996. Pigs experimentally infected with S. hyodysenteriae can be protected from developing swine dysentery by feeding them a highly digestible diet. Epidemiology and Infection, 116, 207-216.
45. TAYLOR, D. J. & ALEXANDER, T. J. L., 1971. The production of dysentery in swine by feeding cultures containing a spirochaete. British Veterinary Journal, 127, 58-61.
46. TAYLOR, D. J., LYSONS, R. J., BEW, J., STEVENSON, R. & LEMCKE, R. M., 1985. Survival of Treponema hyodysenteriae in samples of dysenteric pig faeces sent by post and stored at room temperature. Veterinary Record, 116, 48–49.
47. TROTT, D. J., OXBERRY, S. L. & HAMPSON, D. J., 1997. Evidence for Serpulina hyodysenteriae being recombinant with an epidemic population structure. Microbiology (Reading), 143, 3357–3365.
48. WHITING, R. A., DOYLE, L. P. & SPRAY, R. S., 1921. Swine Dysentery. Purdue University Agricultural Experimental Station Bulletin, 257, 1-15.
49. WINDSOR, R. S. & SIMMONS, J. R., 1981. Investigation into the spread of swine dysentery in 25 herds in East Anglia and assessment of its economic significance in 5 herds. The Veterinary Record, 109, 482–484.
50. WOOD, E. N. & LYSONS, R. J., 1988. The financial benefits from the eradication of swine dysentery. The Veterinary Record, 122, 277–279.
51. ZEEH, F., NATHUES, H., FREY, J., MUELLNER, P. & FELLSTRÖM, C., 2017. A review of methods used for studying the molecular epidemiology of Brachyspira hyodysenteriae. Veterinary Microbiology, 181, 181-194.