Tularemia is a highly contagious disease occurring principally in wild animals but it may transmit to farm animals, causing septicemia and high mortality.
Etiology Francisella tularensis biovar tularensis in North America is tick-transmitted. Francisella tularensis biovar holarctica in Asia, Europe, and North America is transmitted by ticks and mosquitoes
Epidemiology Livestock disease mainly reported from North America; seasonal, associated with heavy tick infestation. Majority of reports in livestock are historical. Zoonosis
Clinical findings Tick infestation. Fever, stiffness of gait, diarrhea, weight loss, recumbency. Wool break
Clinical pathology Culture. Agglutination and intradermal test but not specific
Necropsy findings Subcutaneous swellings at site of tick attachment, lymphadenitis, and septicemia in sheep. Pigs have pleuritis, pneumonia, and abscessation of submaxillary and parotid lymph nodes
Diagnostic confirmation Culture or indirect fluorescent antibody staining
Francisella tularensis is the causative organism. It is Gram-negative, does not form spores and gives partial cross-agglutination with Brucella spp. Isolates are antigenically similar but they can be divided into two types on biochemical and epidemiological features and virulence tests. Type A, Francisella tularensis biovar tularensis, is prevalent in North America. Type B, Francisella tularensis biovar holarctica (palaearctica), is found in Asia, Europe and North America. Type A is associated with tick-borne tularemia in rabbits and type B is associated with mosquitoes and with water-borne disease in aquatic rodents. Type A is the more pathogenic of the two types and the more virulent for humans.1-3 Type B rarely causes disease in higher mammals.
Tularemia is primarily restricted in its occurrence to countries in the northern hemisphere and occurs in most of them. In North America the disease is most prevalent in farm animals in the north-western states of the USA and the adjoining areas of Canada, although in these areas it is rare and the majority of reports in livestock are historical.
F. tularensis has a wide host range and is recorded in over 100 species of bird and wild and domestic animal. Disease is recorded among farm animals, most commonly in sheep and pigs and to a lesser extent in calves, which appear more resistant but can be infected in association with heavy tick infestation.4 Sheep and pigs of all ages are susceptible but most losses occur in lambs, and in pigs clinical illness occurs only in piglets. There is a sharp seasonal incidence, the bulk of cases occurring during the spring months. The morbidity rate in affected flocks of sheep is usually about 20% but may be as high as 40%, and the mortality rate may reach 50%, especially in young animals.
The major reservoirs and transmitters of the infection are rabbits, hares, wild rodents, ticks, and flies and the principal mammalian target host in North America is the cottontail rabbit (Sylvilagus spp.). With sheep, transmission occurs chiefly by the bites of the wood tick, Dermacentor andersoni, and from Haemaphysalis otophila, the ticks becoming infected in the early part of their life cycle when they feed on rodents.5 In Europe Ixodes ricinus and Dermacentor reticulatus are vectors.6
Trans-stadial and transovarial transmission occurs in the tick. The adult ticks infest sheep, and pastures bearing low shrubs and brush are particularly favorable to infestation. The ticks are found in greatest numbers on the sheep around the base of the ears, the top of the neck, the throat, axillae, and udder. It is assumed that sheep are relatively resistant to tularemia but become clinically affected when the infection is massive and continuous. Transmission to pigs and horses is thought to occur chiefly by tick bites but mechanical transmission to laboratory animals does occur with tabanid and blackflies.7
There is little information concerning virulence mechanisms of F. tularensis. The capsule appears to be a necessary component for expression of full virulence and protects against serum-mediated lysis. The lipopolysaccharide has unusual biological and structural properties and low toxicity in vitro and in vivo.3 The organism can persist in dry straw for 6 months, persists for at least 16 months in mud and water and may proliferate in these media.8,9 It does not appear to survive in carcasses for long periods unless they are frozen, when it may persist for 60–120 days.
F. tularensis biovar tularensis has remarkable invasive powers and infection in humans can occur through the unbroken skin. Most exposures appear to result from the handling of infected rabbits and other wildlife but infections can arise from bites of ticks and the deer fly (Chrysops discalis), from the ingestion of contaminated meat and water, and from the bite or scratch of infected cats.10-12 The disease is an occupational hazard to workers in the sheep industry in areas where the disease occurs. Spread of the disease to humans may also occur in abattoir workers who handle infected sheep carcasses. Shearers are susceptible because outbreaks often occur at shearing time. The inhalation or intradermal injection of as few as 10 organisms can establish infection in humans,1,3 and appropriate precautions should be taken in the clinical and postmortem examination of suspect cases. The organism is identified as a potential agent of bioterrorism.13
Tularemia is an acute septicemia but localization occurs, mainly in the parenchymatous organs, with the production of granulomatous lesions.
The incubation period has not been determined. A heavy tick infestation is usually evident.
The onset of the disease is slow with a gradually increasing stiffness of gait, dorsiflexion of the head and a hunching of the hindquarters; affected animals lag behind the group. The pulse and respiratory rates are increased, the temperature is elevated up to 42°C (107°F), and a cough may develop. There is diarrhea, the feces being dark and fetid, and urination occurs frequently with the passage of small amounts of urine. Body weight is lost rapidly, and progressive weakness and recumbency develop after several days, but there is no evidence of paralysis, the animal continuing to struggle while down. Death occurs usually within a few days but a fatal course may be as long as 2 weeks. Animals that recover commonly shed part or all of the fleece but are solidly immune for long periods.
An agglutination test is available for the diagnosis of tularemia, a titer of 1:50 being regarded as a positive test in pigs. Serum from pigs affected with brucellosis does not agglutinate tularemia antigen, but serum from pigs affected with tularemia agglutinates brucellosis antigen. Cross-agglutination between F. tularensis and Brucella abortus is less common in sheep and an accurate diagnosis can be made on serological grounds because of the much greater agglutination that occurs with the homologous organism. Titers of agglutinins in affected sheep range from 1:640 to 1:5000 and may persist at levels of 1:320 for up to 7 months. A titer of 1:200 is classed as positive in sheep. In horses the titers revert to normal levels in 14–21 days.
An intradermal sensitivity test using ‘tularin’ has been suggested as being more reliable as a diagnostic aid in pigs than the agglutination test, but is unreliable in sheep.5
In sheep, large numbers of ticks may be present on the hides of fresh carcasses. In animals that have been dead for some time, dark red subcutaneous areas of congestion up to 3 cm in diameter are found and may be accompanied by local swelling or necrosis of tissues. These lesions mark the attachment sites of ticks. Enlargement and congestion of the lymph nodes draining the sites of heaviest tick infestation are often noted. Pulmonary edema, congestion, or consolidation are inconstant findings.
In pigs the characteristic lesions are pleuritis, pneumonia, and abscessation of submaxillary and parotid lymph nodes. The organisms can be isolated from the lymph nodes and spleen, and from infected ticks. Isolation can also be effected by experimental transmission to guinea pigs. Techniques such as immunoperoxidase staining of fixed specimens and PCR of fresh tissues can circumvent the need for culture of this zoonotic agent.
• Bacteriology – lung, lymph node, spleen (CULT – requires cystine-enriched media, PCR)
• Histology – above tissues plus liver, fixed in formalin (LM, IHC).
Note the zoonotic potential of this organism when handling carcasses and submitting specimens.
The occurrence of a highly fatal septicemia in sheep during spring months when the sheep are heavily infested with Dermacentor andersoni should suggest the possibility of tularemia, especially if the outbreak occurs in an enzootic area.
• Tick paralysis. This occurs in the same area and at the same time of the year as tularemia but is not accompanied by fever and there is marked flaccid paralysis. Recovery from tick paralysis occurs commonly if the ticks are removed
• Other septicemias include P. trehalosi in sheep and Haemophilus spp. in sheep and cattle. These are unusual in the age group in which tularemia occurs and are not associated with tick infestation. In pigs, local lesions can resemble tuberculosis
Streptomycin, gentamicin, the tetracyclines, and chloramphenicol are effective treatments in humans and companion animals.10 Oxytetracycline (6–10 mg/kg BW) has been highly effective in the treatment of lambs and much more effective than penicillin and streptomycin.5
An outbreak of tularemia in sheep can be rapidly halted by spraying or dipping with an insecticide to kill the vector ticks. In areas where ticks are enzootic, sheep should be kept away from shrubby, infested pasture or sprayed regularly during the months when the tick population is greatest. An experimental live attenuated vaccine has been developed but there is no routine vaccination of livestock.
Hopla CE. The ecology of tularaemia. Adv Vet Sci. 1974;18:25-53.
Feldman KA. Tularemia. J Am Vet Med Assoc. 2003;222:725-730.
Tarnvik A, Priebe HS, Grunow R. Tularaemia in Europe: an epidemiological overview. Scand J Infect Dis. 2004;36:350-355.
Petersen JM, Schriefer ME. Tularemia: emergence/re-emergence. Vet Res. 2005;36:455-467.
1 Rohrbach BW. J Am Vet Med Assoc. 1988;193:428.
2 Morner T, et al. J Vet Med B. 1993;40:613.
3 Sjostedt A. Curr Opin Microbiol. 2003;6:66.
4 Philip CB, Williams SC. Bull Soc Vector Ecol. 1985;10:45.
5 Gordon JR. Can J Comp Med. 1983;47:408.
6 Hubalek Z, Halouzka J. J Med Entomol. 1997;34:660.
7 Philip CB, Jellison WL. Bull Soc Vector Ecol. 1986;11:197.
8 Bell JF, et al. Can J Comp Med. 1978;42:310.
9 Feldman KA. J Am Vet Med Assoc. 2003;222:725.
10 Woods JP, et al. J Am Vet Med Assoc. 1998;212:81.
11 Klock LE, et al. J Am Vet Med Assoc. 1973;226:149.
Etiology Yersinia pseudotuberculosis and Yersinia enterocolitica cause occasional disease in farm animals under special circumstances. These organisms also are increasingly recognized as a cause of disease in humans and this is their main importance
Epidemiology Subclinical and clinical disease in ruminants associated with either organism in animals that are debilitated from other influences. Enterocolitis is an emerging form of infection. Pigs carry Y. enterocolitica in the tonsils, pharynx, and intestine but the epidemiology of infection is poorly established
Clinical findings Ruminants: chronic ill-thrift and in a wasted condition with or without diarrhea
There are pathogenic and nonpathogenic strains of Yersinia pseudotuberculosis and Yersinia enterocolitica. The pathogenic strains of both organisms possess chromosomal and plasmid-mediated virulence determinants.1
Y. pseudotuberculosis can be divided into 15 major serogroups, based on O-antigens, some of which can be further divided into subgroups on the basis of type-specific somatic and flagellar antigens. There is variation in animal and human pathogenicity between the serogroups.
Y. enterocolitica is divided into five biovars. It is serologically heterogeneous and 27 serotypes have been identified on the basis of somatic and flagellar antigens with further subdivisions. Bioserotypes may be host-specific. Serotypes O:2, O:3, O:5, O:8, and O:9 have been associated with infection in farm animals and humans, whereas other serotypes appear to be nonpathogenic. Serotype O:9 is antigenically very similar to Brucella spp., and infection with this serotype is a cause of false-positive reactions to Brucella agglutination and complement fixation tests.
Yersiniosis has worldwide occurrence, although there appear to be regional differences in the species of animal infected, the prevalence of disease and the organism involved. Y. pseudotuberculosis has historically been associated with sporadic pyemic disease in sheep manifest with extensive abscessation of internal organs such as liver and spleen. More recently it, and Y. enterocolitica, have been associated with enterocolitis occurring in cattle,2-4 sheep, pigs and goats,3,5 buffalo,6 and farmed and feral deer.7-9 Enteric disease in ruminants has been particularly reported in recent years from Australia and New Zealand.
Y. pseudotuberculosis is a common inhabitant of the intestine in a wide variety of domestic and wild mammals. Wild birds and rodents are also reservoirs of the organism, and fecal–oral spread on pastures and in water is a major method of transmission. Spring migratory birds can spread pathogenic types over long distances.10
There may be differences in the host specificity of different serotypes and strains. Rodents and birds may be the major reservoirs for serotypes I and II, which infect deer and goats,9 whereas sheep and cattle may be a maintenance host for serotype III.
In an Australian study Y. pseudotuberculosis serotype III was isolated from the feces of healthy sheep in 5% of flocks examined and the prevalence was probably much higher as only a small number of sheep were sampled in each flock. Infection was more common in young sheep, occurred during the winter and spring months, and excretion of the organism persisted for 1–14 weeks.2
In cattle the organism has been found without disease in 26% of normal cattle and on 84% of farms tested.11 The fecal excretion that occurs in clinically normal sheep and cattle possibly results from a subclinical infection of the intestine, as experimental challenge of ruminants can result in the establishment of the organism in the intestine with the presence of microscopic abscessation in the lamina propria and serological conversion in the absence of clinical disease.2,5
Enteric disease associated with this organism in both cattle and sheep appears to occur as the result of a heavy infection pressure in animals that are debilitated from other influences. These include cold wet weather, inanition and starvation, trace element deficiency, change of diet, management procedures such as marking and, in farmed deer, procedures such as capture, yarding, and recent transport.2-512
In sheep, attack rates in the flock for clinical disease have ranged from 1–90% with a mean of 18% and a population mortality varying from 0–6.7%.13 Y. pseudotuberculosis may also cause sporadic abortion in cattle and sheep.11,14 In sheep, abortion rates of 1–9% of pregnant sheep are recorded, abortion occurring in the latter part of pregnancy and without clinical illness in the ewes.11 The organism is the cause of occasional cases of bovine caprine mastitis,15,16 epididymitis, and orchitis in rams, and it may be found in sporadic cases of abscessation and lymphangitis in ruminants.
Y. enterocolitica is not commonly associated with clinical disease in farm animals. Diarrhea associated with this organism can occur in sheep, and the organism can be isolated from affected lambs. However pathogenic strains of Y. enterocolitica are less pathogenic to sheep and goats than pathogenic strains of Y. pseudotuberculosis.
Enterocolitis is recorded in sheep and goats.3,13,17,18 Biotype 5, serotype O:2,3 has been isolated from some of these.18 In an Australian survey this organism was detected in 17% of flocks and was isolated from young sheep at all seasons of the year.2 Clinical disease appears to be predisposed by the same stress factors as apply with disease associated with Y. pseudotuberculosis. Disease is recorded in sheep under 1 year of age with an attack rate that varied from 2–55% and a population mortality that ranged from 0.3–16.7%.13,17 Y. enterocolitica is also an occasional cause of abortion in sheep and this has been reproduced experimentally.19
Whereas Y. enterocolitica is commonly isolated from pigs, and pigs are a major reservoir for human disease, it is a rare cause of clinical disease in conventional pigs,20 although clinical enteric disease can be produced by experimental challenge of colostrum-deprived pigs.21 Conventional pigs challenged with serotype 0:3 excreted the organism in feces but were fecal-culture-negative 10 weeks after challenge and at slaughter, even though the organism could be isolated from the tonsils at slaughter. Pigs seroconverted at day 19 postchallenge and remained seropositive until slaughter 70 days later.20
Human infection with Y. pseudotuberculosis is primarily manifest with septicemia, and renal failure is a sequel. In addition to food-borne infection the consumption of water contaminated by animal feces appears to be a major risk factor. Raw milk consumption is also a risk.22
Gastrointestinal disease associated with Y. enterocolitica appears to have increasing prevalence in humans and can be associated with a reactive arthritis as a sequel. Septicemia does occur but is largely limited to those with other underlying disease. Serotypes O:3, O:5, O:8, and O:9 are incriminated. Pigs are a major reservoir for Y. enterocolitica, and pork and pork products are sources for human infection. Serotype O:3, in particular, is commonly isolated from the tonsils and pharynx of pigs at slaughter and less commonly from feces. The rate of isolation varies geographically and with farm source and it has been suggested that pathogen-free breeding is a method for control.23
In contrast to Australia and New Zealand, it is believed that in Europe the pig is the only domestic animal consumed by humans that regularly harbors pathogenic Yersinia.24 There is an apparent increasing prevalence of O:3 infections in humans in the northern hemisphere and pigs and pork products are considered to be important sources.23,25 A survey in Great Britain comparing isolates of Y. enterocolitica from cattle, sheep, and pigs with those from humans over a 2-year period did not find a strong correlation between pathogenic serotypes isolated from the two groups, with the exception of isolates from pigs.26 The importation of meat products has been incriminated as the vehicle of introduction of pathogenic serotypes into Japan.27 There would appear to be an increased risk for infection in humans handling pigs at slaughter and in veterinarians in pig practice.
Invasion of the intestinal epithelium is followed by inflammation in the mucosa and the formation of microabscesses in the lamina propria and mesenteric lymph nodes. Ulcers and disruption of the intestinal mucosa lead to loss of fluid and function. The intestinal lesions are accompanied by villous atrophy and lead to malabsorption and ill-thrift, diarrhea or a combination of the two.13
Affected animals may present with a syndrome of chronic ill-thrift and in a wasted condition with or without diarrhea. Where diarrhea is present the feces are watery, foul-smelling, and black in color, but occasionally they also contain mucus and blood. Diarrhea persists for 2–3 weeks in an individual animal and may require yarding and dagging as control procedures to avoid fly strike.
There is a neutrophilia with a left shift. Affected animals are often hypoproteinemic and anemic, although this may be a reflection of the underlying malnutrition. In experimental infections antibody develops by 9–19 days postinfection and may be an aid to diagnosis.5,20 The organism can be isolated from the feces. A multiplex PCR is capable of detecting 10 pathogenic serobiotypes of Y. enterocolitica28 and PCR has been developed to discriminate pathogenic Y. enterocolitica from other members of this genus.29
There are liquid intestinal contents but usually no gross findings. Some sheep may have thickening of the mucosa of the small intestine and the cecum and colon, and the mesenteric lymph nodes may be enlarged and edematous.
The characteristic findings on histopathology consist of a segmental suppurative erosive enterocolitis.13 Microabscesses, consisting of aggregations of neutrophils with prominent colonies of Gram-negative coccobacilli, are present in the mucosa. Lesions are most prevalent in the jejunum and ileum and are accompanied by atrophy of villi and hyperplasia of cryptal epithelium. Microabscesses may coalesce to produce extensive erosions and there may be microabscesses in the liver.
The placenta from sheep that have aborted in association with Y. pseudotuberculosis is thickened and edematous with necrotic debris in the intercotyledonary zone and must be differentiated from enzootic abortion.14
• Bacteriology – jejunum, ileum, colon, mesenteric lymph node (CULT – sometimes requires cold enrichment)
• Histology – formalin-fixed jejunum, ileum (several sections), colon, mesenteric node (LM).
Note the zoonotic potential of this organism when handling carcasses and submitting specimens.
Isolates vary in their sensitivity to antibiotics and a sensitivity test is advisable. Most isolates show in vitro sensitivity to the aminoglycosides, to tetracyclines and to sulfonamides or a combination of sulfonamides and trimethoprim.13,23,30 Sulfonamides and trimethoprim are reported not to be effective in the treatment of yersiniosis in cattle; long-acting tetracyclines are recommended for the treatment of both infections, in combination with supportive therapy.31
There is no specific control but the maintenance of good nutrition is believed to be an important factor in avoiding clinical disease.
1 Nagano T, et al. J Vet Med Sci. 1997;59:153.
2 Slee KJ, Skilbeck NW. J Clin Microbiol. 1992;30:712.
3 Gill J. Surveillance. 1996;23:24.
4 Slee KJ, et al. Aust Vet J. 1988;65:271.
5 Slee KL, Button C. Aust Vet J. 1990;67:320.
6 Hum S, et al. Aust Vet J. 1997;75:95.
7 Sanford SE. J Vet Diagn Invest. 1995;7:78.
8 Wilson PR. Surveillance. 2002;29:13.
9 Jerett IV, et al. Aust Vet J. 1990;67:212.
10 Niskanen T, et al. Appl Environ Microbiol. 2003;69:4670.
11 Hodges RT, Carman MG. N Z Vet J. 1985;33:175.
12 McLennan MW, Kerr DR. Aust Vet J. 2000;78:28.
13 Philbey AW, et al. Aust Vet J. 1991;68:108.
14 Otter A. Vet Rec. 1996;138:143.
15 Jones TO, et al. Vet Rec. 1982;112:231.
16 Bleul U, et al. Vet Rec. 2002;151:767.
17 McSporran KD, et al. N Z Vet J. 1984;32:38.
18 Slee KL, Button C. Aust Vet J. 1990;67:396.
19 Corbel MJ, et al. Br Vet J. 1992;148:339.
20 Nielsen B, et al. Vet Microbiol. 1996;48:293.
21 Shu D, et al. N Z Vet J. 1995;43:50.
22 Bahout AA, Moustafa AH. Assuit Vet Med J. 2004;50:57.
23 Hariharan H, et al. Can J Vet Res. 1995;59:161.
24 Nesbaklen T, et al. Contrib Microbiol Immunol. 1991;12:32.
25 Thibodeau V, et al. Vet Microbiol. 2001;82:249.
26 McNally A, et al. Lett Appl Microbiol. 2004;39:103.
27 Fukushima H, et al. J Food Microbiol. 1998;35:205.
28 Harnett N, et al. Epidemiol Infect. 1996;117:59.
29 Arnold T, et al. Syst Appl Microbiol. 2001;24:285.
Atrophic rhinitis is a disease affecting primarily young pigs but causing anatomical lesions which may persist for life. The term nonprogressive atrophic rhinitis is used for the slight to severe rhinitis and usually transient turbinate atrophy in which no toxigenic P. multocida are found, when there are no clinical signs and no obvious growth retardation. This mild form is probably as a result of infection with Bordetella bronchiseptica and/or nontoxigenic P. multocida. The term progressive atrophic rhinitis is proposed for the infection with toxigenic P. multocida (capsular serotype D and A strains) characterized by shortening or distortion of the snout, sneezing, nasal discharge, and epistaxis. Progressive atrophic rhinitis is often accompanied by reduced growth rates in severe cases.
Etiology Toxigenic strains of Bordetella bronchiseptica and Past. multocida
Epidemiology Young growing pigs. High percentage of pigs reared under intensive conditions may have some degree of atrophic rhinitis. Infection widespread and transmitted by carrier sow to piglet. Housing and ventilation risk factors. Immunity develops in herd. Major economic importance because may affect growth rate and predispose to pneumonia
Signs Initially sneezing when piglets 3–9 weeks of age. Nasal discharge. Deformity of face with nasal bones (twisted snout). Growth rate may be decreased
Clinical pathology Culture organism from nasal swabs. PCR
Lesions Varying degrees of severity of atrophic rhinitis
Diagnostic confirmation Necropsy examinations of snouts
Treatment Antimicrobials in early stages; nothing later
Control Eliminating toxigenic strains of Past. multocida. Depopulation and repopulation. Reduction of infection. Mass medication. Medicated early weaning. Vaccination
Infection of the nasal cavities with B. bronchiseptica followed by toxigenic strains of P. multocida – primarily capsular type D1-4 and occasionally type A – results in progressive turbinate atrophy.5 P. multocida type A strains were formerly thought to be associated entirely with lung infections but there is increasing evidence that some strains of P. multocida type A are increasingly toxin producers and may be involved in atrophic rhinitis.6,7 Toxin production appears to be independent of serotype. The strains of P. multocida isolated from the lungs are usually nontoxigenic and of capsular type A8 but a small proportion are toxigenic and/or possess capsular type D.8,9 A complex study of the porcine strains from progressive atrophic rhinitis and pneumonia has been completed.10 Eighteen groups of strains were identified on the basis of specific combinations of capsular type, tox A status and outer membrane protein (OMP) type. The majority (88%) of cases of pneumonia were associated exclusively with nontoxigenic capsular type A strains of OMP types 1.1, 2.1, 3.1, and 5.1 and capsular type D isolates of OMP6.1. They are primary pathogens with a relatively high degree of virulence.10 In contrast the majority (76%) of cases of progressive atrophic rhinitis were associated with tox A-containing capsular type D strains of OMP4.1 and capsular type A and D strains of OMP6.1. They further found that toxigenic capsular type A strains associated with progressive atrophic rhinitis and nontoxigenic strains of capsular type A associated with pneumonia represented separate populations of P. multocida recognizable by their OMP types.10 The results of this analysis led the authors10 to conclude that horizontal transfer of capsular biosynthesis and tox A genes has occurred between certain strains of P. multocida.
Atrophic rhinitis occurs worldwide where pigs are reared under intensive conditions. It has, however, become much less important with the onset of vaccination, improvement in resistance by pig breeding companies and general attention to the environment in the farrowing house. It is one of the most common diseases in pigs in the midwestern USA. Some surveys have shown that 50% of finished pigs and sows at slaughter have lesions of atrophic rhinitis. The incidence of clinical disease varies from 5–30%, which in part depends on the method of detection of the gross lesions. Abattoir surveys of the snouts of slaughtered pigs indicate that the incidence of gross lesions ranges from 14–50%. However, the incidence of gross lesions in abattoir surveys is biased by the source of the pigs; the incidence may be low in pigs from herds that have attempted to control the disease and high in some commercial herds with no control program. In pigs slaughtered from pig testing stations the incidence of lesions may be uniform over a long period.3 The published data on the incidence of gross lesions are also variable because of the lack of a uniform method of evaluating and quantifying the lesions. However, major improvements in the development of uniform and repeatable methods of evaluating lesions have occurred in recent years.
The incidence and severity of the lesions may vary with the season and the type of facility in which pigs are reared. In a slaughter survey of the snouts and lungs of pigs from 21 pig herds over one winter and one summer the lesions of atrophic rhinitis were more severe among pigs slaughtered in the summer, whereas lesions of pneumonia were more severe among pigs slaughtered in the winter.11 Lesions of atrophic rhinitis were also more severe in pigs farrowed in central, enclosed farrowing houses and finished in enclosed, mechanically ventilated buildings than in pigs farrowed individually in sow huts and finished on dirt lots. It is possible that the incidence and severity of the lesions at slaughter may be a reflection of the condition of the housing facilities when the animals were piglets several months previously, but many other factors could have been involved.
B. bronchiseptica readily colonizes the ciliated mucosa of the respiratory tract of pigs and infection of the nasal cavities of pigs is present in almost every pig herd, with the prevalence of infection in pigs in commercial herds varying from 25–50%. Serological surveys of individual herds have found that up to 90% of the pigs are positive, which indicates that there is no reliable correlation between the frequency of isolation of the organism and the percentage of animals with antibody. The prevalence of infection is just as high in specific-pathogen-free herds as in non-specific-pathogen-free herds.
The prevalence of infection of toxigenic P. multocida type D is higher in herds with clinical disease. The organism can be present in 50–80% of weaned pigs in a herd with clinical disease in the finishing pigs. Toxigenic type D P. multocida was first detected in New South Wales, Australia, in 1986; in all herds examined, the introduction of pigs from an infected herd in South Australia was associated with an increased risk of infection.12 Toxigenic P. multocida type D has been isolated rarely from herds free of atrophic rhinitis.13
Whereas B. bronchiseptica is eliminated from the respiratory tract of most infected pigs, leaving only a few infected at slaughter, P. multocida often persists.
Direct contact and droplet infection are presumed to be the most likely methods of transmission. The reservoir of infection is the infected sow, and litters of piglets become infected at an early age. Colonization of the tonsil by P. multocida in conventionally reared pigs is common.14 In the Netherlands it has been recognized that infection is usually by one of four possibilities.15 These are artificial insemination centers, laborers, neighborhood infection by direct aerosol or indirect local contact, and the presence of carrier animals and birds.
The infection is usually introduced into a herd by the purchase of infected pigs. Spread between piglets is probably enhanced after weaning when mixing of litters occurs and 70–80% of a large weaned group become infected. Infection persists for up to several weeks and months, followed by a gradual reduction in the intensity and rate of infection. In herds where B. bronchiseptica is the initiating agent, up to 90% of pigs 4–10 weeks of age will have nasal infection, but this infection rate falls to approximately 15% by 12 months of age and the proportion of carrier pigs within the breeding herd decreases with increasing age of sow. The prevalence of infection is also much higher during the period from October to March than at other times of the year, and the prevalence of serologically positive animals is highest from July to December. This is most probably a result of the winter housing conditions, with few air changes per hour, fluctuating temperatures, and high humidity.
The epidemiology of toxigenic strains of P. multocida as a causative agent of atrophic rhinitis is not as well understood. The organism colonizes the tonsils of clinically normal pigs. In contrast to B. bronchiseptica, which is ubiquitous in pig herds the toxigenic isolates of P. multocida appear to be restricted to herds affected with progressive atrophic rhinitis.5 The organism is invariably present in herds with progressive atrophic rhinitis but may also be present in about 5% of the pigs in a herd with no clinical history of atrophic rhinitis. The main source of toxigenic isolates of P. multocida for young pigs appears to be the pharyngeal tissues of the breeding stock. About 10–15% of sows in farrowing houses may be infected with toxigenic isolates and piglets become infected within a week after birth. In contrast to B. bronchiseptica, infection of piglets at 12–16 weeks of age with toxigenic P. multocida will still result in varying degrees of severity of lesions.
It is possible for growing pigs to develop lesions of atrophic rhinitis well beyond the age of 3 weeks if they are exposed to pigs affected with disease and infected with P. multocida and B. bronchiseptica.
The age at which piglets first become infected with B. bronchiseptica has an important effect on the development of lesions. The most severe lesions occur in nonimmune animals infected during the first week of life. Animals infected at 4 weeks of age develop less severe lesions, while those infected at 10 weeks do not develop significant lesions.
The level of immunity in the young pigs will influence the level of infection and the incidence of clinical disease. Colostral immunity from sows serologically positive to B. bronchiseptica is transferred to piglets and provides protection for 2–5 weeks. Clinical disease does not occur in piglets with high levels of passive antibody. Older pigs from 10–12 weeks of age may become infected but are less likely to develop severe turbinate atrophy and may develop inapparent infection and become carriers.
Vaccination of the sow before parturition to increase colostral immunity or vaccination of the young pig will increase the rate of clearance of the organism from the nasal cavity and reduce the incidence of clinical disease. In chronically affected herds a level of immunity develops with increasing age of the breeding herd.
The virulence characteristics of B. bronchiseptica and the toxigenic isolates of P. multocida are important risk factors. Both organisms are required to produce lesions similar to the naturally occurring progressive disease. The virulence of B. bronchiseptica is dependent on the ability to produce heavy, persistent colonization in the nasal cavity and the production of a heat-labile toxin. Bordetellas produce several virulence factors and toxins, which are regulated by a two-component sensory transduction system encoded by the bvg locus.16,17 These virulence factors include adhesins such as filamentous agglutinin, pertactin, and fimbriae, and the adenylate cyclase–hemolysin toxin and the dermonecrotic toxin. In cell cultures the dermonecrotic toxin stimulates DNA and protein synthesis and assembly of actin stress fibers while inhibiting cell division, resulting in polynucleation of cells.18,19 It mediates these through the modification and activation of the small guanosine 5′-triphosphate (GTP)-binding protein Rho.20,21
There are both toxigenic and nontoxigenic strains of B. bronchiseptica.22 Colonization of the nose was greater with the dermonecrotic-toxin-positive strains than with the dermonecrotic-toxin-negative mutant strains.23 This was maintained for the first, second, and third weeks postinoculation but by the fourth week the position had changed to the opposite. All dermonecrotic-toxin-positive pigs had pneumonia but the dermonecrotic-toxin-negative animals were able to colonize the lung more freely. There is an outer membrane protein P68 perlactin (B. bronchiseptica perlactin gene (prn)) that is an adhesin, which may play a part in the protective immunity24 and may be extremely variable.25 The most important experiment is one that shows that P. multocida mutant strains without the capacity to produce P. multocida type D toxin did not produce turbinate atrophy.26 Only certain porcine phase 1 cultures possess both properties. However, even the most virulent of 10 isolates of B. bronchiseptica did not cause progressive turbinate atrophy or significant snout deformation in experimental infections. The severe lesions of atrophic rhinitis cannot be attributed to this organism alone. Experimental inoculation of specific-pathogen-free or gnotobiotic pigs with the organism results in a nonprogressive moderately severe turbinate atrophy 2–4 weeks after infection, followed frequently by regeneration of the turbinates. These virulence characteristics of B. bronchiseptica are consistent with the observations that in herds where the organism is common it can provoke sneezing and coughing but no evidence of clinical turbinate atrophy. Examination of the turbinates within 2 weeks after the sneezing will reveal some mild lesions but no lesions will be evident when the pigs are examined at slaughter. It may be that the adhesins left over in the nasal cavity from an infection of B. bronchiseptica are subsequently available for the attachment of other bacteria.27
Toxigenic isolates of P. multocida colonize the nasal cavities, elaborate several toxins, and produce progressive lesions of the turbinate bones and snout. Toxigenic P. multocida can colonize the upper respiratory tract of pigs28 and the presence of the capsule is a virulence factor.29 The presence of B. bronchiseptica can enhance the colonization of P. multocida, particularly the toxigenic type D strains isolated from pigs. The cytotoxin of B. bronchiseptica is required for optimum growth by toxigenic P. multocida; other products of phase 1 B. bronchiseptica growth assist colonization by P. multocida, and the degree of atrophy of the turbinates in these mixed infections is related to the numbers of toxigenic P. multocida in the nasal cavity.30 Severe turbinate damage and shortening of the snout can be reproduced in specific-pathogen-free and gnotobiotic pigs by combined infection with B. bronchiseptica and certain strains of P. multocida. Following experimental infection both organisms may persist in the nasal cavities for up to 64 days. The cell envelope proteins and lipopolysaccharides of P. multocida strains associated with atrophic rhinitis have been characterized and compared. At least three protein patterns and six lipopolysaccharide patterns can be distinguished, which can be used to predict the pathogenic character of some of the strains. This will obviate the need to use the guinea-pig skin test to distinguish those strains that are associated with atrophic rhinitis and those that are not.
The gene for the osteolytic toxin of P. multocida has been cloned and expressed in E. coli; the protein expressed has been shown to have the same properties as the native toxin. The toxin is the main colonization factor produced by toxigenic strains of the organism and antitoxin made from the toxin is protective experimentally and cross-protective between toxins from different capsule types.31 The toxin can produce turbinate atrophy when injected intranasally and also when given intramuscularly, intraperitoneally, intravenously, or intradermally.32 Fingerprinting techniques have been used to show that outbreaks of atrophic rhinitis since 1985 in Australia have been associated primarily with a single strain of toxigenic type D P. multocida.12
The effects of housing, population density and adequacy of ventilation on the prevalence of infection of B. bronchiseptica and toxigenic isolates of P. multocida, and on the incidence and severity of atrophic rhinitis, have not been examined in detail. Atmospheric ammonia, dust, and microbial concentrations in the farrowing house and dust in weaner barns have a significant role in the severity of atrophic rhinitis.33,34 The mean daily gain of gilts with atrophic rhinitis exposed to ammonia may be smaller than that of those not affected.35 Undocumented field observations suggest that the disease is more common and severe when pigs are confined, overcrowded and housed in poorly ventilated unsanitary barns, all of which promote the spread of infection.
There is no effect of high levels of ammonia on the severity of turbinate atrophy. It has been shown that high levels of ammonia have no effect on the disease progression of atrophic rhinitis and pneumonia but do enhance the colonization of the nasal turbinates by toxigenic P. multocida.36 A recent experiment has shown34 that higher numbers of P. multocida bacteria were isolated from the tonsil than the nasal membranes per gram of tissue. Aerial pollutants contribute to the severity of lesions associated with atrophic rhinitis by facilitating colonization of the upper respiratory tract by P. multocida.
Management factors such as confinement farrowing and the use of continual throughput farrowing houses and weaner houses are also considered to be important risk factors. Adverse climatic conditions (below thermoneutrality with drafty periods) can result in a lower amount of energy available for production because of increased maintenance requirements,37 which results in growth retardation associated with lowered feed intake.
Historically, it was accepted as dogma that atrophic rhinitis was an important cause of economic loss in pig herds because of decreased growth rate, less than optimal feed efficiency and the fact that it was a major risk factor in enzootic swine pneumonia. A number of field studies have found an association between atrophic rhinitis and reduced growth rate in some herds, whereas other observations were unable to show an association between the presence of the disease and growth rate. The lack of a standard system for evaluation of conchal lesions may be a factor in the variable results between observations.38
Some field studies have failed to show that the disease has an effect on growth rate in finishing pigs or that there is a cause and effect relationship between atrophic rhinitis and pneumonia. The presence of pneumonia in pigs from a test station reduced mean daily weight gains by 33% for each 10% of affected lung, but atrophic rhinitis did not affect daily gain and there was no association between the development of atrophic rhinitis and the development of pneumonia. Pigs vaccinated against B. bronchiseptica had turbinate atrophy scores or mean daily gains no different from those of unvaccinated pigs. In another study there was a low positive correlation between the herd mean turbinate atrophy score and the herd mean percentage pneumonia score. A recent report from Illinois indicates that the prevalence of clinical atrophic rhinitis in farrow-to-finish herds ranged from 0–20% and in pigs from those herds examined at the abattoir the incidence of turbinate lesions ranged from 5–92%. In some of the herds the mean daily weight gain was 15–18% higher than in herds where pigs had severe turbinate lesions. In an Australian report there was no correlation between the severity of atrophic rhinitis and growth rate or back fat thickness.
In one study of three commercial pig herds, the snouts and lungs of individual pigs were examined and scored at slaughter and the results were correlated with growth indicators for each pig (average daily gain during the growing and finishing phases, and days to reach market). Scores for lung lesions were also correlated to scores for snout lesions. Contrary to findings in many other studies, pigs that reached market weight at the youngest age did not have the lowest score for lung lesions, nor the lowest grade for snout lesions, nor the least extensive or severe lesions.39 It was concluded that lung lesions and grades for snout lesions in pigs at slaughter are not valid indicators for determining the economic effect of either pneumonia or atrophic rhinitis on growth performance of pigs.
Following infection of the nasal cavity, B. bronchiseptica becomes closely associated with the ciliated epithelium of the respiratory tract. The organism produces a heat-labile toxin that results in a nonprogressive, moderately severe turbinate atrophy that is apparent within 2–4 weeks after infection, followed frequently by regeneration of the conchae. There is, initially, ciliary loss and ciliary stasis, followed by reduction in mucociliary clearance followed by hyperplasia and metaplasia of the nasal epithelium, fibrosis in the lamina propria, and resorption and replacement fibrosis of the osseous core. Experimental infection with B. bronchiseptica alone does not result in severe persistent conchal atrophy or twisting or shortening of the snout. The strains of B. bronchiseptica that produce cytotoxin may predispose to the colonization of P. multocida in the nasal cavities.
Infection and colonization of the nasal cavities with the toxigenic strains of P. multocida results in the elaboration of a toxin that causes progressive conchal atrophy. The toxin is thermolabile and dermonecrotic and is called the dermonecrotic toxin of P. multocida. The inoculation of a toxin from a toxigenic strain of type D P. multocida into the nasal cavities of gnotobiotic pigs results in severe bilateral atrophy of the conchae. Atrophy of the ventral conchae can be produced experimentally with pathogenic B. bronchiseptica in piglets at 6 weeks of age and with toxigenic P. multocida strains in piglets as old as 16 weeks of age.
The toxin enhances osteoclastic resorption and impairs osteoblastic synthesis of the conchal osseous core; irreversible changes can occur within a few days.40 The toxin is able to subvert cell cycle progression and cell–cell signaling systems in osteoblasts and osteoclasts.41,42 The toxin is the sole agent responsible for the conchal atrophy and the effect appears to be related to the total exposure to the toxin, i.e. dose-dependent.43 More importantly, this also appears to have an immunomodulatory effect. There is an inverse relationship between the number of P. multocida and the total concentration of immunoglobulin.44 This may in part be one of the reasons that local changes in the nose produce such adverse growth effects, and they may be due to the fact that the P. multocida type D toxin has in fact changed the immune functions and that the P. multocida may have predisposed to many other agents.43 These authors’ conclusion is that P. multocida significantly suppresses the antigen-specific IgG immune responses of pigs to parenteral antigen challenge.43 The epithelium and the submucosa undergo secondary atrophy and the conchae may disappear almost completely within 10–14 days. These lesions can persist until the animal is 90 kg in body weight. The conchal atrophy is not accompanied by an inflammatory reaction. The effect of the P. multocida toxin is restricted to the nasal cavity; this is supported by the intriguing observation that the parenteral injection of the toxin into gnotobiotic piglets results in turbinate lesions and shortening and twisting of the snout. The parenteral injection of the dermonecrotoxin of P. multocida capsular type D into specific-pathogen-free adult pigs will result in moderate conchal atrophy.45 In piglets 7 days of age, the intramuscular injection of the purified dermonecrotoxin will result in severe atrophy of the conchae.45 The culture filtrate of a non-atrophic-rhinitis pathogenic P. multocida will not cause lesions after intramuscular injection. The disappearance of the conchae and the involvement of the bones of the face lead to deformity of the facial bones with the appearance of dishing and bulging of the face and, if the lesion is unilateral, to lateral deviation of the snout.
The effect on growth rate, if any, may be due to the chronic irritation and interference with prehension. Experimentally, atrophic rhinitis suppressed the health of pigs, reducing their activity and feed intake.46 Experimentally, parenteral injections of the toxin decrease physeal area and reduce chondrocyte proliferation in long bones, in addition to conchal atrophy.47
Reliable experimental models of atrophic rhinitis in gnotobiotic pigs are now available and are useful for studying the pathogenesis of the disease and testing vaccine strategies. A sterile sonicate of a toxigenic strain of B. bronchiseptica is instilled into the nasal cavities of piglets at 5 days of age followed by intranasal inoculation of toxigenic strains of P. multocida at 7 days of age.48
The clinical findings of atrophic rhinitis depend on the stage of the lesions. In acute cases in piglets 3–9 weeks of age, irritation of the nasal mucosa causes sneezing, some coughing, small amounts of serous or mucopurulent nasal discharge, and transient unilateral or bilateral epistaxis. The frequency of sneezing may be a measure of the incidence and severity of the disease. In piglets born from sows vaccinated with B. bronchiseptica and P. multocida vaccine before farrowing, followed by two vaccinations within 3 weeks of age, the frequency of sneezing at 3–9 weeks of age was much less than in piglets given only B. bronchiseptica vaccine. There may be rubbing of the nose against objects or on the ground. A watery ocular discharge usually accompanies this and may result in the appearance of dried streaks of dirt below the medial canthus of the eyes. There may be a decrease in growth rate. In infection with B. bronchiseptica these clinical signs will disappear spontaneously in a few weeks, when the pigs will appear normal. In severe cases, respiratory obstruction may increase to the point of dyspnea and cyanosis, and sucking pigs may have great difficulty in nursing. The nasal secretions become thicker and nasal bleeding may also occur.
In the more chronic stages, inspissated material may be expelled during paroxysms of sneezing. During this chronic stage, there is often pronounced deformity of the face due to arrested development of the bones, especially the conchae, and the accumulation of necrotic material in the nasal cavities. The nasal bones and premaxillae turn upwards and interfere with approximation of the incisor and, to a lesser extent, the molar teeth. There are varying degrees of brachygnathia superior and protrusion of the lower incisor teeth. Prehension and mastication become difficult, with a resulting loss of body condition. Facial distortion in the final stages takes the form of severe ‘dishing’ of the face with wrinkling of the overlying skin. If the condition is unilateral, the upper jaw may be twisted to one side. These visible facial deformities develop most commonly in pigs 8–10 weeks old within 3–4 weeks after infection, but they may occur in younger pigs.
The most serious effects of the advanced disease are depression of growth rate and unthriftiness. The appetite may be unaffected but much feed is lost by spillage and feed efficiency may be reduced in some instances.
It is important to be able to detect infected animals in a herd, especially the carrier animal. Nasal swabs are used to detect the bacteria and to determine their drug sensitivity. The collection of the nasal swabs must be done carefully and requires a special transport medium to insure a high recovery rate. A sampling technique and a special culture medium to facilitate the isolation and recognition of B. bronchiseptica are described.49 The external nares are cleaned with alcohol and a cotton-tipped flexible wire is pushed into the nasal cavity (of each side in turn) until it reaches a point midway between the nostril and the level of the medial canthus of the eye. On removal, the cotton tip is cut off into 0.5 mL of an ice-cold sterile transport medium comprising phosphate-buffered saline (PBS, pH 7.3) with fetal calf serum (5% v/v). The samples are then placed on special media, preferably within 4 hours. Normally the organism grows well on conventional culture media, especially when younger pigs are sampled. However, in the carrier pig the organism may be sparse and the selective medium is recommended.
The nasal culturing procedure has been used as an aid in the control of atrophic rhinitis associated with B. bronchiseptica. A series of three nasal swabs from each animal is considered to be about 77% efficient in detecting infected animals for possible culling and elimination from the herd. However, in some studies there may be no marked difference in the prevalence of B. bronchiseptica or P. multocida in pig herds with or without clinical atrophic rhinitis.49
Toxigenic P. multocida grow readily in the laboratory but are difficult to isolate from nasal swabs because they are frequently overgrown by commensal flora. Selective laboratory media containing antimicrobial agents have been developed to promote the isolation of P. multocida from nasal swabs. Inoculation of cotton swabs to selective medium on the same day as the sampling provides the best isolation of toxigenic P. multocida. Immersion of pigs at slaughter in the scalding tank can result in a marked reduction in the isolation of toxigenic P. multocida.
A cell culture assay using embryonic bovine lung cell cultures is available and is a sensitive in vitro test for the differentiation of toxigenic from nontoxigenic isolates of P. multocida. This test can replace the lethal tests in mice or the dermonecrotic tests in guinea-pigs.
Agglutination tests and an ELISA test are available for the detection of pigs infected with B. bronchiseptica, especially carrier animals. Serology is of value in the assessment of the response of pigs vaccinated with the B. bronchiseptica vaccines. There are currently no reliable serological tests for Pasteurella.
A PCR method originally described in 199650 for the enhanced detection of toxigenic P. multocida directly from nasal swabs has been described and upgraded.51 This was shown to be 10 times more sensitive than P. multocida type D toxin (PMT) ELISA and five times more sensitive than clinical bacteriology with subsequent use of PMT ELISA. A nested PCR has also been described.52 Similarly, a PCR method for the detection of B. bronchiseptica has been described53 that produces 78% more positives than culture, particularly with swabs with a high mixed bacterial load. Recently a nested-PCR has been described that was reported to be more specific and sensitive than the other PCR methods previously described.54 It does not require culture, it is less laborious and the results can be provided within 24 hours. The authors concluded that this test was suitable for breeding company evaluations and for eradication schemes.
The typical lesions of atrophic rhinitis are restricted to the nasal cavities, although concurrent diseases, especially virus pneumonia of pigs, may produce lesions elsewhere. In the early stages there is acute inflammation, sometimes with the accumulation of pus, but in the later stages, there is evidence only of atrophy of the mucosa, and decalcification and atrophy of the conchae and ethmoid bones, which may have completely disappeared in severe cases. The inflammatory and atrophic processes may extend to involve the facial sinuses. There is no evidence of interference with the vascular supply to the affected bones. The changes in the nasal cavities are most readily seen if the head is split in the sagittal plane but for accurate diagnosis the degree of conchal symmetry, volume and atrophy and medial septum deviation should be assessed by inspection of a vertical cross-section of the skull made at the level of the second premolar tooth.
The clinical diagnosis is confirmed and the severity of the lesions is assessed by the postmortem examination of a cross-section of the snout. The snout must be sectioned at the level of the second premolar tooth because the size of the conchal bone reduces anteriorly and may give a false-positive result if the section is taken too far forward. Quantification of the severity of the lesions has been of value for monitoring the incidence and severity of the disease in a herd. Several systems have been used for grading the severity of lesions of the snout. Most of them have used a subjective visual scoring system in which snouts are grade 0 (complete normality) to 5 (complete conchal atrophy). Reasonable agreement among observers recording morphological changes of nasal conchae is achievable with some training.49
The standards for each grade are as follows:55
Grade 0: No deviation from absolute normality, with nasal septum straight and conchae symmetrical and filling nasal cavities
Grade 1: Slight irregularity, asymmetry or distortion of the nasal structures without atrophy
Grade 2: Marked distortion of nasal structure but without marked atrophy
Grade 3: Definite atrophy of the conchae with or without distortion
Grade 4: More severe atrophy with severe atrophy of one or more conchae
Grade 5: Very severe atrophy in which all conchae have virtually disappeared.
Such a discontinuous grading system does not provide a direct quantitative relationship. Regular examination of the snouts from heads of pigs sent to slaughter can be used to assess the level of conchal atrophy in the herd. Morphometric methods, using either point counting or semi-automated planimetry applied to photographic or impression prints of sections of the snout to measure the extent of conchal atrophy on a continuous scale as a morphometric index, are now available. Cross-sections of the snout are photographed or used to make impression prints, which are then measured.56 A morphometric index is determined, which is the ratio of free space to total cross-sectional area of the nasal cavity. The system correlates well with the visual grading system of 0–5 but is labor-intensive and relatively expensive. The conchal perimeter ratio may be a more reliable morphometric measure of atrophic rhinitis and also provides parametric data suitable for quantitative analysis.57 A morphometric analysis using conchal area ratio is the best method for quantifying gross morphological turbinate changes.58 Descriptions of the methods for making snout impressions are available.59,60 Computed tomography has been described.61
A major limitation of the grading system is that conchal atrophy occurs as a continuous spectrum and it is difficult to decide, for example, if a pig with a grade 3 lesion represents the more severe manifestation of B. bronchiseptica infection, which may not progress further, or an early manifestation of infection with toxigenic P. multocida, which could develop into a severe herd problem.
The occurrence of sneezing in the early stages and of facial deformity in the later stages are characteristic of this disease.
Inclusion body rhinitis due to a cytomegalovirus is a common infection in young piglets in which there is sneezing and conjunctivitis. However, by itself it does not progress to produce turbinate atrophy and facial distortion. Under good hygienic conditions the course of the disease is about 2 weeks and the economic effects are minimal. In the early acute stages, atrophic rhinitis may be mistaken for swine influenza which, however, usually occurs as an outbreak affecting older pigs and accompanied by a severe systemic reaction without subsequent involvement of facial bones.
Necrotic rhinitis is manifested by external lesions affecting the face, and virus pneumonia of pigs is characterized by coughing rather than sneezing.
The inherited prognathic jaw of some breeds of pigs has been mistaken for the chronic stage of atrophic rhinitis; protrusion of the lower jaw is quite common in adult intensively housed pigs and has been attributed to behavioral problems of pushing the snout against fixed equipment such as bars and nipple drinkers.
Treatment early in the course of the disease will reduce the severity of its effects, but it is of little value in chronically affected pigs, and these pigs are best culled at an early age because of their persistent poor growth rate and high food conversion.
Tylosin at 20 mg/kg BW, oxytetracycline at 20 mg/kg BW, or trimethoprim– sulfadoxine (40 mg/200 mg/mL) at 0.1 mL/kg BW may be given parenterally or the creep feed may be medicated with sulfamethazine and/or tylosin at 200 and 100 mg/kg of feed respectively. Parenteral injections need to be repeated every 3–7 days for at least three injections and feed medication should be given for 3–5 weeks. The problem with early creep medication is in obtaining adequate intakes of the antibacterial. This is seldom achieved before 2 weeks of age and parenteral antibiotics may be required if significant infection occurs before this stage.
The parenteral administration of antimicrobial agents to individual piglets at 3–7-day intervals beginning at 3 days of age for a total of three to five injections per piglet has been recommended for the treatment and control of atrophic rhinitis. However, in a large herd such a treatment regimen would be a major task and, until a cost–benefit analysis indicates a beneficial effect over other methods, we cannot recommend such a practice.
The treatment of experimental B. bronchiseptica infection in young pigs has been successful with the use of trimethoprim–sulfadiazine in the drinking water at levels of 13.3 and 77.6 μg/mL respectively, for 3 weeks. This method would remove the necessity to inject pigs repeatedly.
Tilmicosin has proved useful;62,63 fed continuously over 6 weeks at concentrations of 200 g per ton of feed it controlled transmission of atrophic rhinitis, weight gains were positively affected, and fewer nasal swabs were positive for P. multocida at the end of the study period.64
Effective control depends on developing methods of eliminating or controlling the prevalence of toxigenic isolates of P. multocida, which cause progressive atrophic rhinitis if they become established in the nasal cavity. Previous infection of the nasal cavity with B. bronchiseptica can enhance the establishment of toxigenic P. multocida and result in progressive atrophic rhinitis.
While there is considerable information available on the ecology of B. bronchiseptica and the methods by which it might be eliminated or controlled in a herd, there is little documented information available on methods that can be used for control of the toxigenic isolates of P. multocida associated with atrophic rhinitis.
Control of atrophic rhinitis can be attempted in at least four ways:
• Reduction of infection pressure
• Mass medication with antimicrobials to reduce the severity and adverse effects of infection
Regardless of the method employed, any effective control program must have a system for monitoring the incidence of clinical disease in the herd and the incidence and severity of conchal lesions of the pigs sent to slaughter. Accurate and reliable methods for monitoring clinical disease are not available but the incidence of acute rhinitis and facial deformities could be recorded regularly. At slaughter, snouts can be examined for lesions of conchal atrophy and for assessing a mean snout score for each group of pigs slaughtered.
Total eradication can only be achieved with confidence by complete depopulation for a 4-week period and repopulation with primary or purchased specific-pathogen-free stock. This approach has the added advantage of also eliminating enzootic pneumonia, which may be a significant contributing factor to the economic importance of this disease. However, this method of control is extremely costly and the economic importance of the disease would need to be carefully evaluated in relation to this cost before this method was instituted. Other techniques of obtaining pigs free of atrophic rhinitis, such as the isolated farrowing of older and presumed noncarrier sows with subsequent clinical and postmortem examinations of a proportion of the litters, have had a significant failure rate in the field and are not recommended. Eradication by repopulation with cesarean-derived stock may be essential in breeding nucleus herds where a high generation turnover results in a low herd sow age and a low herd level of immunity. The breakdown rate of herds established by this method can be significant, presumably because the initiating organisms are not solely confined to pigs.
A pilot control scheme was initiated in Britain in which a herd had to meet the following conditions:
• It must be inspected by a veterinarian every 6 months over a period of 2 years, over which time there must be no clinical evidence of atrophic rhinitis
• The herd owner must certify that atrophic rhinitis has not been suspected over the same time period
• Cross-sections of snouts taken from at least 30% of marketed pigs must be examined regularly by a veterinarian, and over a 2-year probationary period the average 6-monthly snout score must not exceed 0.5
• There must be no vaccination or treatment for atrophic rhinitis
• New breeding stock can be introduced only from other qualified herds or herds derived by hysterectomy, artificial insemination, or embryo transfer techniques.
Over a 5-year period 45 herds qualified at some stage, and 34 were still qualified at the end of 5 years. As of 1988, some herds had exceeded the snout score limit of 0.5, with their average scores increasing to 2.24.65 In these herds, there was no clinical, epidemiological, or bacteriological evidence that they were at risk of developing severe atrophic rhinitis. It is suggested that the higher scores were associated with a group of recurrent husbandry factors, especially overstocking and unsatisfactory conditions in the weaner barns. These increased scores suggested the possibility that the upper limit for the snout scores in qualifying herds could be raised and allow bacteriological testing to be confined to more doubtful herds.
Eradication in the Netherlands was based on the fact that they thought that there were four main possibilities for the spread of toxigenic P. multocida: artificial insemination centers, laborers, neighborhood infection either by aerosol or by local spread, and carrier animals or birds. They assumed that most herds were closed or buying certified stock and that the major source of infection was therefore the boar.26 In this study they tested boars; in herds with less than 50 boars they tested all and in those with more than 50 they tested 50 as the minimum. They took nasal and tonsil samples,66,67 which were placed in cold transport medium68 and sent to the laboratory within 24 hours under cooled conditions for overnight culture followed by PCR.69
Reduction of infection pressure can be attempted. Infection of piglets occurs primarily either from carrier sows or from other infected piglets in the immediate environment and severe atrophic rhinitis generally results from infection of piglets under 3 weeks of age. If these factors can be minimized the incidence and severity of the disease can be reduced.70 An all-in/all-out pig flow is one of the most effective methods of control of atrophic rhinitis. Changing to an all-in/all-out pig flow from continuous flow management can improve snout scores by 50%, lung scores by 55%, average daily gain by 0.14 lb and days to market by 13 days.71
Since severe lesions depend upon infection of the piglet under 3 weeks of age, every attempt should be made to minimize the severity of the challenge to young piglets. It is a common observation that the effects of atrophic rhinitis are minimal under good systems of management and adequate ventilation, nondusty conditions and good hygiene. The use of continual-throughput farrowing houses and weaner houses allows a buildup of infection with the presence of actively infected pigs that can provide a high infection pressure on piglets born into or introduced into these areas. The use of all-in/all-out systems of management in these areas is recommended and young piglets should be kept in a separate area from older pigs.
The prophylactic use of antimicrobials is frequently employed to reduce the incidence of the disease within the herd. Antimicrobials are used both within the breeding herd to reduce the prevalence of carriers and in young suckling and weaner pigs to reduce the severity of the infection. The medication is begun about 2 weeks before farrowing, continued throughout lactation and incorporated in the creep feed for the sucking pigs and the starter feeds for the weaned pigs. In this way there is continuous medication of the sow and the piglets during the most susceptible period. For the breeding herd, sulfamethazine at levels of 450–1000 mg/kg feed, with the higher levels being given to dry sows on restricted feeding, has been recommended. Sulfonamide resistance has proved a problem in some countries but beneficial results may still be achieved with these levels. It is recommended that medication be continued for a 4–6-week period. Carbadox at a level of 55 ppm in combination with sulfamethazine at 110 ppm is reported to be effective in clearing experimentally induced B. bronchiseptica infection, and when used alone improved growth rate and feed efficiency in pigs with naturally occurring atrophic rhinitis. In the starter period, carbadox fed alone or in combination with sulfamethazine improved average daily gain in piglets from herds with naturally occurring atrophic rhinitis.72 Use of the medication, however, did not result in a reduction of mean nasal lesion scores due to atrophic rhinitis. Sulfamethazine at 110 mg/kg of feed is more effective than sulfathiazole at the same concentration for the control of experimentally induced atrophic rhinitis due to B. bronchiseptica. Sulfamethazine may also be incorporated in creep rations and the use of tetracyclines (200 mg/kg), tylosin (50–100 mg/kg), and penicillin (200 mg/kg) have also been suggested.
Medicated early weaning is recommended to obtain pigs free from pathogens, including B. bronchiseptica, that are endemic in the herd of origin. The sows are fed medicated feed from 5 days before to 5 days after weaning and the piglets are dosed from birth to 10 days of age.
There has been considerable interest in the development of vaccines for the control and prevention of atrophic rhinitis due to B. bronchiseptica. Inactivated vaccines have been used to vaccinate the pregnant sow 4–6 weeks before farrowing, and in some cases, followed by vaccination of the piglets at 7 and 28 days of age. In general, the use of the vaccine in pregnant sows in herds where the disease has been endemic has reduced the incidence of clinical atrophic rhinitis. However, the results from one study to another have been highly variable. Vaccination of the pregnant sow results in an increase in colostral antibody titer, which does improve the clearance rate of B. bronchiseptica in the piglets. However, it has been difficult to evaluate the efficacy of the B. bronchiseptica used alone because the turbinate atrophy associated with infection of piglets with B. bronchiseptica experimentally or naturally heals and regenerates completely when they are reared to about 70–90 kg BW in good housing conditions.
Vaccination with both components (B. bronchiseptica and P. multocida) in a vaccine reduces lesions considerably when compared with a placebo and a group with only P. multocida type D toxin in the vaccine73 but neither vaccine eliminated toxigenic P. multocida from the upper respiratory tract.
Experimentally, piglets born from sows vaccinated with P. multocida are protected from a challenge with atrophic rhinitis toxin. This indicates that artificial immunization for atrophic rhinitis should be possible. Vaccination of sows at least three times before farrowing for the first time and during each subsequent pregnancy with a vaccine containing B. bronchiseptica and P. multocida was highly successful in reducing the incidence of atrophic rhinitis in the pigs. The incidence in affected herds was reduced from 7.5% to about 2%. Experimentally, the vaccine provides good protection against challenge in piglets from vaccinated sows.72
A recombinant P. multocida toxin derivative vaccine given to gilts 4–5 weeks before farrowing and again 2–3 weeks later provided excellent protection in their piglets against experimental challenge with B. bronchiseptica and toxigenic P. multocida.74 This indicates the excellent immunoprotective properties of the nontoxic derivative of the P. multocida toxin. In five field trials a single component vaccine containing a nontoxic but highly immunogenic protein, the d0-protein, as the antigen, provided much better protection than the control vaccine containing killed P. multocida and killed B. bronchiseptica.75
Experimental infection and vaccination of pregnant minimum-disease sows with B. bronchiseptica resulted in much higher agglutinins in serum and colostrum than in sows only vaccinated or control animals, and the piglets were provided with protection against experimental disease. Vaccination of pregnant gilts with purified inactivated P. multocida toxin resulted in a high degree of protection of their progeny against progressive atrophic rhinitis.76
A new vaccine has been described77 using a truncated P. multocida type D toxin that is immunogenic and nontoxic, a toxoid for B. bronchiseptica and an adjuvant. Sows were vaccinated at 8–6 weeks and 4–2 weeks before farrowing. The vaccinated animals had fewer organisms.
Chanter N, Jones PW, Barnard S, Pennings A. Atrophic rhinitis of swine. In: Manual of standards for diagnostic tests and vaccines, list A and B: diseases of mammals, birds and bees. Paris: Office International des Epizoöties; 2000:615-622.
Magyar T, Lax AJ. Atrophic rhinitis. In: Brogden KA, Gunthmiller JM, editors. Polymicrobial diseases. Washington, DC: ASM Press; 2002:169-197.
1 Eamens GJ, et al. Aust Vet J. 1988;65:120.
2 Foged NT, et al. J Clin Microbiol. 1988;26:1419.
3 Gardner IA, et al. J Vet Diagn Invest. 1994;6:442.
4 Lariviere S, et al. J Clin Microbiol. 1992;30:1398.
5 Chanter N. Pig News Info. 1990;11:503.
6 Fussing V, et al. Vet Microbiol. 1999;65:61.
7 Sakano T, et al. J Vet Med Sci. 1992;54:403.
8 Rubies X, et al. Vet Microbiol. 2002;84:69.
9 Choi C, et al. Vet Rec. 2001;149:210.
10 Davies RL, et al. J Med Microbiol. 2003;52:59.
11 Cowart RP, et al. J Am Vet Med Assoc. 1991;200:190.
12 Gardner IA, et al. Aust Vet J. 1991;68:364.
13 Kavanaugh NT, et al. Vet Rec. 1994;134:218.
14 Ackermann MR, et al. J Vet Diagn Invest. 1994;6:375.
15 De Jong M, et al. In: Proceedings of the 17th International Pig Veterinary Society Congress 2002:329.
16 Arico B, et al. Mol Microbiol. 1991;5:2481.
17 Monack DM, et al. Mol Microbiol. 1989;3:1719.
18 Horoguchi Y, et al. Infect Immun. 1993;61:3611.
19 Horoguchi Y, et al. FEMS Microbiol Lett. 1994;106:19.
20 Horoguchi Y, et al. FEMS Immunol Med Microbiol. 1995;12:29.
21 Horoguchi Y, et al. Proc Natl Acad Sci U S A. 1997;94:11623.
22 Magyar T, et al. Vet Microbiol. 1988;18:135.
23 Brockmeier SL, et al. Vet Microbiol. 2000;73:1.
24 Novotny P, et al. Infect Immun. 2000;50:190.
25 Register KB. In: Proceedings of the 16th International Pig Veterinary Society Congress 2000:483.
26 Brockmeier SL, et al. Infect Immun. 2002;70:481.
27 Tuomanen E. Infect Immun. 1986;54:905.
28 Pijoan C, Trigo F. Am J Vet Res. 1990;54:516.
29 Jacques M, et al. Infect Immun. 1993;61:4785.
30 Chanter N, et al. Res Vet Sci. 1989;47:48.
31 Lax AJ, Chanter N. J Gen Microbiol. 1990;136:81.
32 Williams PP, et al. Can J Vet Res. 1990;54:157.
33 Robertson JF, et al. Anim Prod. 1990;50:173.
34 Hamilton TDC, et al. Clin Diagn Lab Immunol. 1999;6:199.
35 Diekman MA, et al. Am J Vet Res. 1993;54:2128.
36 Andreasen M, et al. J Vet Med. 2000;47:161.
37 Van Diemen PM, et al. Livestock Prod Sci. 1995;43:275.
38 Cowart RP, et al. J Am Vet Med Assoc. 1990;196:1062.
39 Scheidt AB, et al. J Am Vet Med Assoc. 1990;196:881.
40 Ghoshal NG, Niyo Y. Am J Vet Res. 1994;54:738.
41 Rozengurt E, et al. Proc Natl Acad Sci U S A. 1990;87:123.
42 Mullan PB, Lax AJ. Infect Immun. 1996;64:959.
43 Jordan RW, et al. FEMS Immunol Med Microbiol. 2003;39:51.
44 Elias B, et al. J Vet Med Sci. 1996;55:617.
45 Martineau-Doize B, et al. Can J Vet Res. 1991;55:224.
46 Van Diemen PM, et al. J Anim Sci. 1995;73:1658.
47 Ackermann MR, et al. Am J Vet Res. 1996;57:848.
48 Ackermann MR, et al. Infect Immun. 1991;59:3626.
49 Lariviere S, et al. J Clin Microbiol. 1993;31:364.
50 Kamp EM, et al. J Vet Diagn Invest. 1996;8:304.
51 Ohlinger VF, et al. In: Proceedings of the 16th International Pig Veterinary Society Congress 2000:478.
52 Choi CS, Chae CH. Vet J. 2001;162:255.
53 Reizenstein E, et al. Diagn Microbiol Infect Dis. 1993;17:185.
54 Sulko C, et al. In: Proceedings of the 17th International Pig Veterinary Society Congress 2002:90.
55 Jackson GH, et al. Br Vet J. 1982;138:480.
56 Done JT, et al. Vet Rec. 1984;114:33.
57 Collins MT, et al. Am J Vet Res. 1989;50:42.
58 Gatlin CL, et al. Can J Vet Res. 1996;60:121.
59 Done JT, et al. Br Vet J. 1984;140:418.
60 Lund LJ, Beckett P. Vet Rec. 1983;113:473.
61 Magyar T, et al. Acta Vet Hung. 2003;51:485.
62 Clark LK, et al. J Swine Health Prod. 1998;6:257.
63 Olson LB, Backstrom LR. J Swine Health Prod. 2000;8:263.
64 Backstrom L, et al. J Swine Health Prod. 1994;2:11.
65 Goodwin RFW. Vet Rec. 1988;123:566.
66 De Jong MF, et al. In: Proceedings of the 13th International Pig Veterinary Society Congress 1994:238.
67 De Jong MF, et al. In: Proceedings of the 14th International Pig Veterinary Society Congress 1996:249.
68 De Jong MF, et al. In: Proceedings of the 16th International Pig Veterinary Society Congress 2000:477.
69 Baekbo P, et al. In: Proceedings of the 13th International Pig Veterinary Society Congress 1988:51.
70 Pejsak Z, et al. Compend Immunol Microbiol Infect Dis. 1994;17:125.
71 McCaw MB. Compend Contin Educ Pract Vet. 1994;16:1615.
72 Kobisch M, Pennings A. Vet Rec. 1989;124:57.
73 Magyar T, et al. In: Proceedings of the 16th International Pig Veterinary Society Congress 2000:479.
74 Nielsen JP, et al. Can J Vet Res. 1991;55:128.
75 Bording A, et al. Acta Vet Scand. 1994;35:155.
Diseases Associated With Brucella Species
The species of Brucella and their principal farm animal hosts are Brucella abortus (cattle), Brucella melitensis (goats), Brucella suis (pigs), and Brucella ovis (sheep). In general, the principal manifestations of brucellosis are reproductive failure, such as abortion or birth of unthrifty newborn in the female, and orchitis and epididymitis with frequent sterility in the male. Persistent (lifelong) infection is a characteristic of this facultative intracellular organism, with shedding in reproductive and mammary secretions. Brucellosis is also an important zoonosis causing debilitating disease in humans. Because of the major economic impact on animal health and the risk of human disease, most countries have attempted to provide the resources to eradicate the disease from the domestic animal population.
Control programs have employed two principal methods: vaccination of young or mature animals and the slaughter of infected and exposed animals, usually on the basis of a reaction to a serological test.
Brucellosis has been eradicated from cattle in several regions of the world and is nearly eradicated in others. However, it is still widespread and is an economically important agricultural disease in many countries. There are still many cases of human brucellosis reported each year in regions where the disease has not been eliminated in farm livestock.
In 2002, an entire issue of the journal Veterinary Microbiology, vol 90, pp 1–603, was devoted to brucellosis. Several review articles on various aspects of brucellosis from that special issue are listed below under Review literature.
The history of brucellosis is fascinating.1 In 1884, Captain David Bruce and several others working on Mediterranean fever isolated an agent they called Micrococcus melitensis from human spleens. Hospital patients were fed raw goat’s milk for many illnesses and this was an early example of a nosocomial infection. The ship SS Joshua Nicholson, when anchored in Malta in 1905, took on board 65 goats bound for Washington. The Bureau of Animal Industry of the USDA had decided to import Maltese goats to encourage goat husbandry among peasant immigrants from Southern Europe. Nearly all the ship’s crew drank the raw milk and, within weeks, were ill. The goats were never shipped to the USA, on the recommendation of Bruce. Goat’s milk was banned from the military garrison in 1906, essentially ending the problem among naval forces, but the controversy about goats persisted for many years. The goat had a special niche in Maltese culture and was taken door to door providing fresh milk. A pasteurization commission was formed in 1938 and but not enforced until 1959. Even as late as 1955, over 200 human cases of brucellosis were associated with ingestion of a special cheese. Human brucellosis exists worldwide and is spread mainly by ingestion of unpasteurized dairy products. An overview of the literature on human brucellosis is available.2
In 1985, Professor L.F. Benhard Bang, Danish veterinary pathologist and bacteriologist, described a different causative organism isolated from cattle, called Bacillus abortus. The first recognized human case of brucellosis in the USA was in 1898 in an army officer who contracted the disease in Puerto Rico. In the early 20th century contagious abortion of cattle and tuberculosis were recognized as major causes of economic loss. By 1922, several states had passed laws and regulations in an attempt to prevent introduction of the disease in cattle purchased from other states. In 1930, concerns were raised about the relationship between the disease in animals and humans. The American Veterinary Medical Association recommended a field trial of a vaccine, which was developed from a strain of lower virulence named Brucella abortus strain 19. This vaccine has been used several decades as the most common immunizing agent for control of bovine brucellosis.
In 1934, a cooperative State Federal Brucellosis Eradication program was launched on a nationwide scale in the USA. A uniform plan provided for blood tests, slaughter of seropositive cattle and federal indemnities. In 1941, strain 19 was introduced and used in most states. All vaccinated cattle had to be properly identified. In 1934/35, the reactor rate in cattle tested was 11.5%. In 1954, the US Congress appropriated funds for a comprehensive national effort to eradicate brucellosis. The brucellosis eradication program was designed as a cooperative effort between the federal government, the states, and livestock producers. In 1957, shortly after the inception of the program, there were almost 124 000 infected herds identified. As of December 2004, there were no brucellosis-affected cattle herds in the USA; truly a success story in veterinary medicine.3 When brucellosis can be identified, contained and eliminated before spread occurs, eradication can be achieved.
The evolution of the Brucella spp. and its taxonomy have been described in detail.4 In 2002, the genomes for B. melitensis were published, and B. suis and B. abortus are in their final stages.5 The proteomes of selected Brucella spp. have been extensively analyzed, all of which will provide opportunities to understand the complete biology of different Brucella species.6 The major outer membrane proteins of Brucella spp. have also been determined.7 The lipopolysaccharide of Brucella is unique and considered as a virulence factor that helps bacterial survival by circumventing the immune response.8 The pathology of brucellosis reflects the outcome of a battle between the host genome and the Brucella genome.9
In addition to brucellosis occurring in terrestrial wildlife, Brucella spp. have been isolated from some marine mammals of North America10 and there is serological evidence of Brucella spp. infection from several species of marine mammal from both hemispheres.11 Serological evidence of Brucella spp. has been found in odontoceles (suborder of toothed whales, including sperm whales, porpoises, grampuses, dolphins, beaked whales, bottle-nosed whales, and narwhals) from the south Pacific and the Mediterranean.11
The status of brucellosis in regions of the world and countries varies considerably. The following is a summary of the status of brucellosis in various regions of the world as of 2002.
Romania, like many other developed countries, eradicated B. abortus from cattle in 1969.12 The incidence of brucellosis in sheep and pigs is rare and B. melitensis has never been reported. Vaccination against brucellosis is prohibited.
In Macedonia and Greece, brucellosis occurs in sheep, goats, and humans, associated with B. melitensis.13 In Greece, cows are infected with B. abortus or B. melitensis. In Croatia, B. suis biovar 2 is found in pigs. In Yugoslavia, brucellosis is endemic in some regions. A financially well-supported control and eradication program such as that sponsored by the European Union is needed.
The overall serological prevalence of infection is 6.26% in sheep, 7.24% in goats and 0.58% in cattle.14 B. melitensis predominates as the cause of brucellosis in ruminants in Kosovo.
Brucellosis is an important disease among livestock and humans in sub-Saharan Africa.15 The disease in cattle is prevalent and widespread and is caused primarily by B. abortus; B. melitensis and B. suis have been suspected. In sheep and goats, B. melitensis is common and the prevalence of infection is high. Brucellosis has occurred in pigs in these countries but information is limited. Brucellosis due to B. abortus is one of the most important diseases of camels in the arid and semi-arid pastoralist areas of central, east, and west Africa.
Seroepidemiological surveys for brucellosis in Eritrea found a prevalence of 8.2% in dairy cattle, with a herd prevalence of 35.9%.16,17 The prevalence in sheep and goats was variable depending on the geographical area. In camels, the seroprevalence was 3.1%.
In the near East region, animal brucellosis affects almost all domestic animals, particularly cattle, sheep, and goats.18 Brucellosis occurs in camels in Saudi Arabia, Kuwait, Oman, Iraq, Iran, Sudan, Egypt, Libya, and Somalia. In Egypt, brucellosis occurs in cattle, buffaloes, horses, and pigs. B. melitensis biovar 3 is the most commonly isolated species from animals in Egypt, Jordan, Israel, Tunisia, and Turkey. The highest incidence of human brucellosis occurs in Saudi Arabia, Iran, Palestinian Authority, Syria, Jordan, and Oman. Bahrain has none. Most human cases are associated with B. melitensis, biovar 3 but B. abortus is increasingly being reported in humans. The control of brucellosis in these countries is very controversial, with varying emphases on different aspects of control. The most commonly used vaccines are B. abortus strain 19, B. melitensis Rev. 1, and B. abortus RB 51.
B. abortus is a major cause of abortion among cattle and buffaloes.19 The incidence of the disease is low and the small size of the country would facilitate an effective disease control program.
Brucellosis was first recognized in India in 1942 and is endemic throughout the country.20 The disease occurs in cattle, buffalo, sheep, goats, pigs, dogs, and humans. B. abortus biotype 1 in cattle and buffaloes and B. melitensis biotype 1 in sheep, goats, and humans are the predominant infective biotypes. Economic losses are considerable in an agrarian country such as India. There is no organized and effective brucellosis control program. Plans for a large-scale control program, including calfhood vaccination, are underway.
A major constraint of a control program is that all slaughter of cows is banned and that segregation of seropositive cows until their death will therefore be necessary, but very costly.
Before the 1980s, human and animal brucellosis was severe. B. melitensis is most common in outbreaks.21 Sheep, cattle, and pigs are the main sources of infection for humans. Beginning in 1950, control programs have been in place and progress is being made towards control and eradication.
Bovine brucellosis due to B. abortus is the most prevalent Brucella infection in Brazil, followed by B. suis in pigs. B. melitensis and Brucella neotomae have not been isolated. The prevalence of bovine brucellosis ranged from 4–5% in the period of 1989–1998. The disease is considered endemic, with a higher incidence in regions with a higher cattle density.22 In 2001, a New National Program was launched, including compulsory vaccination of heifers aged 3–8 months, voluntary accreditation of free herds, voluntary monitoring of beef herds based on periodic sampling, regulatory tests for breeding stock prior to interstate movement and entrance into livestock fairs and exhibitions, compulsory slaughter of cattle testing positive, and standardization of testing procedures through short courses for accredited veterinarians.
Brucellosis has existed in the country for many years. Most reports are on B. abortus in cattle, but B. melitensis and B. suis have been identified. In 2000, it was estimated that the prevalence of B. abortus in the cattle population was 3.15%.23 A national campaign for the control and eradication of brucellosis was begun in 1978. The program is based on vaccination of calves at 3–8 months of age, testing and culling of seropositive animals, declaration of Brucella-free areas, and promotion of Brucella-free Herd Certification Programs in dairy herds. For beef herds there is mandatory vaccination, control of movement of animals intended for breeding, including testing for those imported, destined to fairs and auctions.
Brucellosis continues to be a serious disease for animal and human health in Venezuela.24 B. abortus is the most common biovar, causing high rates of abortion in cattle and buffalo. Based on the rapid agglutination plate test, the positive reactor rate ranges from 0.8–1.2%; using the ELISA the prevalence is 10.5%. A control program, in effect since 1968, consists of vaccination of calves with strain 19 vaccine, and test and slaughter of positive reactors. Improved testing methods and vaccination of all female calves between 3 and 8 months of age, and revaccination at 10–15 months of age and adult cattle in high prevalence areas.
B. abortus and B. suis infections occur in all Central American countries, and sheep and goat brucellosis associated with B. melitensis occurs in Guatemala.25 The estimated prevalence of bovine brucellosis ranges from 4–8% with a herd prevalence (dairy herds) of 10–25%.25
A national control program based on vaccination of calves and test and slaughter of reactors has been unsuccessful. Possible reasons include inadequate economical support for vaccination and test and slaughter programs, and the high density of Brucella infections.
Brucellosis has been recognized in Argentina since the 19th century.26 In 2000, the individual cow prevalence was 5% and the herd prevalence 10–15%.26 In dairy cattle, the prevalence is estimated at 2–2.5%. The annual economic losses due to the disease in cattle have been estimated at US$60 000 000. A control program began in 1932 and successive changes have been issued since then. The current program mandates vaccination of all females with B. abortus strain 19 between 3 and 8 months of age, and test and slaughter of positive animals. However, the compensation paid for reactors is inadequate and producers commonly retain the reactors in the herd. The program has been most successful in dairy herds, which receive incentive payments if the prevalence of infection is low. The disease has been found in pigs, goats, sheep, and dogs. Human brucellosis is an important disease in Argentina. Federal financial support is needed to assist the livestock industry to eradicate the disease.
Brucellosis is an important disease in Mexico.27 Five of the seven known Brucella species have been isolated, including B. melitensis biovars 1–3; B. abortus biovars 1, 2, 4–6; B. suis biovar 1; Brucella canis and B. ovis. Brucellosis is endemic in the cattle, sheep, and goat populations. The disease is a trade barrier. Each year Mexico exports 1.2 million steers and heifers to the USA. The heifers must be spayed to minimize the risk of brucellosis transmission. A control program has been in effect since 1942 but vaccination was voluntary and the disposition of reactors inconsistent. Brucellosis is a significant public health problem in Mexico because 35% of the milk and cheeses consumed are unpasteurized. As of 2002, about 3500 cases of human brucellosis are reported annually and it is estimated this figure represents only one-third of the actual cases. About 98% of cases are due to ingestion of contaminated dairy products (mainly goat cheeses). About 93% of human cases are infected with B. melitensis of goat origin. In 1993, a control program was reinforced with the creation of the National Commission for Bovine Tuberculosis and Brucellosis Eradication. In high-risk areas, massive vaccination programs of goats with B. melitensis Rev. 1 strain in adults and young females are being implemented, along with a Sanitary Package to improve goat and sheep health. In 1997, the use of B. abortus RB 51 vaccine was officially approved. Mexico is one of the few countries authorized to produce this live attenuated vaccine. To reduce abortions, a reduced dose is commercially produced for adult females. As of 2000, almost 1 million beef and dairy cattle, and 1 million goats are vaccinated annually; almost 97% of the dairy cattle population is vaccinated.
The status of bovine brucellosis in Canada and the USA is presented below under Bovine brucellosis associated with Brucella abortus.
Austria, Denmark, Finland, Germany, the Province Bolanzo (Italy), Luxembourg, Norway, Sweden, the Netherlands, and Great Britain have gained the status of being officially brucellosis free.28 Countries not officially brucellosis-free are France, Greece, Ireland, Italy, Portugal, and Spain. The prevalence of infection in countries not free of brucellosis is extremely diverse. The highest numbers of infected herds occurred in southern Europe: Greece, Spain, Italy, and, Portugal. The prevalence of infection has increased in both the Republic of Ireland and Northern Ireland.
Both B. abortus and B. melitensis have been isolated from cattle; B. melitensis may be isolated from cattle in contact with sheep and goats. B. melitensis may cause isolated cases of abortion in cattle rather than outbreaks of abortion.
Great Britain has been free from brucellosis since 1993 and is required by the European Union regulations to test 20% or more of both beef and dairy cattle below 24 months of age routinely.
Belgium, Denmark, Finland, Germany, the Netherlands, Ireland, Austria, Luxembourg, Norway, Sweden, the UK, Spain, and France are officially free of ovine and caprine brucellosis. Greece, Italy, and Portugal are not officially free. Ovine and caprine brucellosis is a significant problem for both public health and animal production in Greece, and a control program consists of vaccination of lambs and kids, and test and slaughter policy. European co-financed eradication programs for sheep and goat brucellosis are in place in France, Greece, Italy, Portugal, and Spain.28
Brucellosis in pigs, especially outdoor-raised pigs, has re-emerged as a result of infection from the wild boar brucellosis (B. suis biovar 2) reservoir. In all EU countries and Norway, boars are subject to pre-entry testing and regular testing every 18 months at artificial insemination stations.
Adams LG. The pathology of brucellosis reflects the outcome of the battle between the host genome and the Brucella genome. Vet Microbiol. 2002;90:553-561.
Cloeckaert A, Vizcaino N, Paquet J-Y, et al. Major outer membrane proteins of Brucella spp. past, present and future. Vet Microbiol. 2002;90:229-247.
DelVecchio VG, Wagner MA, Eshenbrenner M, et al. Brucella proteomes: a review. Vet Microbiol. 2002;90:593-603.
Letesson J-J, Lestrate P, Delrue R-M, et al. Fun stories about Brucella: the ‘furtive nasty bug’. Vet Microbiol. 2002;90:317-328.
Michaux-Charachon S, Jumas-Bilak E, Allardet-Servent A, et al. The Brucella genome at the beginning of the post-genomic era. Vet Microbiol. 2002;90:581-585.
Moreno E, Cloeckaert A, Moriyon I. Brucella evolution and taxonomy. Vet Microbiol. 2002;90:209-227.
Nicoletti P. A short history of brucellosis. Vet Microbiol. 2002;90:5-9.
Ragan VE. The Animal and Plant Health Inspection Service (APHIS) brucellosis eradication program in the United States. Vet Microbiol. 2002;90:11-18.
Doganay M, Aygen B. Human brucellosis: an overview. Int J Infect Dis. 2003;7:173-182.
Lapaque N, Moriyon I, Moreno E, Gorvel J-P. Brucella lipopolysaccharide acts as a virulence factor. Curr Opin Microbiol. 2005;8:60-66.
1 Nicoletti P. Vet Microbiol. 2002;90:5.
2 Doganay M, Aygen B. Am J Infect Dis. 2003;7:173.
3 Ragan VE. Vet Microbiol. 2002;90:11.
4 Moreno E, et al. Vet Microbiol. 2002;90:209.
5 Michaux-Charachon S, et al. Vet Microbiol. 2002;90:581.
6 DelVecchio VG, et al. Vet Microbiol. 2002;90:593.
7 Cloeckaert A, et al. Vet Microbiol. 2002;90:229.
8 Lapaque N, Moriyon I, Moreno E, Gorvel J-P. Curr Opin Microbiol. 2005;8:60-66.
9 Adams LG. Vet Microbiol. 2002;90:553.
10 Nielsen O, et al. J Wildl Dis. 2001;37:89.
11 Van Bressem M-F, et al. Vet Rec. 2001;148:657.
12 Dobrean V, et al. Vet Microbiol. 2002;90:157.
13 Taleski V, et al. Vet Microbiol. 2002;90:147.
14 Jackson R, et al. Vet Rec. 2004;154:747.
15 McDermott JJ, Arimi SM. Vet Microbiol. 2002;90:111.
16 Omer MK, et al. Vet Microbiol. 2002;90:257.
17 Omer MK, et al. Epidemiol Infect. 2000;125:447.
18 Refai M. Vet Microbiol. 2002;90:81.
19 Bandara AB, Mahipala MB. Vet Microbiol. 2002;90:197.
20 Renukaradhya GJ, et al. Vet Microbiol. 2002;90:183.
21 Dequi S, et al. Vet Microbiol. 2002;90:165.
22 Poester FP, et al. Vet Microbiol. 2002;90:55.
23 Baumgarten D. Vet Microbiol. 2002;90:63.
24 Francisco J, Vargas O. Vet Microbiol. 2002;90:39.
25 Moreno E. Vet Microbiol. 2002;90:31.
26 Samartino LE. Vet Microbiol. 2002;90:71.
27 Luna-Martinez JE, Mejia-Teran C. Vet Microbiol. 2002;90:19.