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Haemophilus parasuis infection

Haemophilus parasuis infection

M-L PENRITH AND M M HENTON

GENERAL INTRODUCTION: FACULTATIVELY ANAEROBIC GRAM-NEGATIVE RODS Haemophilus and Histophilus spp. infections

Introduction

Haemophilus parasuis typically causes Glässer’s disease, a peracute to acute disease of 2- to 16-week-old pigs characterized by a serofibrinous polyserositis, polyarthritis and meningitis. In some cases, acute septicaemia results in death before typical lesions develop.1, 3, 47 In conventional herds, occurrence is sporadic and morbidity and mortality rates are generally low, although outbreaks may occur. During the present decade, H. parasuis has emerged as a cause of serious disease in specific pathogen-free (SPF) pigs.35 In South Africa, Glässer’s disease is relatively rare, but increasing interest in and importation of SPF pigs could alter the present situation.

A disease syndrome manifested by fibrinous pleuritis, pericarditis and peritonitis, and often accompanied by arthritis, was described by Glässer in 1910.9 He was able to demonstrate Gram-negative bacilli in the exudate but could not cultivate them. Some decades later it was observed that meningitis was often an associated lesion in animals suffering from the syndrome.13 In 1939, a bacterium thought to be Haemophilus suis, which had first been described in 1931 in association with lesions caused by swine influenza,41 was isolated from the joints of pigs suffering from arthritis.40 This was followed some years later by the isolation of the same bacterium from typical cases of Glässer’s disease and by the experimental production of the disease in pigs by inoculation of H. suis. 13

In 1969, a new species of Haemophilus (H. parasuis) isolated from pigs was proposed.4 This organism required V-factor or nicotinamide adenine dinucleotide (NAD) for growth, but not X-factor (protoporphyrin IX or protohaeme). During recent years this has been the most frequent bacterium isolated from suspected cases of Glässer’s disease.37 Most strains in culture collections identified as H. suis require only V-factor and therefore belong to H. parasuis, 21 because H. suis requires both X- and V-factors. Haemophilus parasuis is therefore considered to be the only cause of Glässer’s disease.

Aetiology

Bacteria of the genus Haemophilus (meaning blood-loving) belong to the family Pasteurellaceae. Haemophilus parasuis (meaning suis-like) is a small, thin, pleomorphic, Gramnegative rod.4 The bacterium is V-factor dependent and therefore requires the addition of this factor to the culture medium. Growth on chocolate agar is feeble after 48 to 72 hours; in the absence of the V-factor, the colonies are smooth, greyish and translucent, and reach a diameter of about 0,5 mm. There is, however, marked enhancement of growth on blood agar if the medium is streaked with a Staphylococcus feeder culture (as staphylococci, as well as certain other bacteria, produce the critical growth factor in excess) with colonies attaining a diameter of 1 to 2 mm around the streak.21 Chocolate agar and Levinthal media (agar and broth) are satisfactory media for propagation of the bacterium.21Growth is also improved if the carbon dioxide level is raised to about 5 per cent.

Haemophilus parasuis is not haemolytic, ferments glucose, sucrose, maltose, galactose, mannose, fructose, ribose and insulin, and is urease- and Camp-negative.21 Identification of H. parasuis is based on morphological and colonial characteristics, V-factor dependency and biochemical testing. For this purpose reference should be made to Bergey’s Manual of Systematic Bacteriology. 22

As has been reported, 44 Bakos, Nilsson and Thal in 1952 proposed a serological classification of H. parasuis based on a tube agglutination test of whole cell preparations. Isolates of groups A, B or C are serologically related organisms; all other strains are placed in an ill-defined group D. In an attempt to provide a more comprehensive system, Morozumi and Nicolet in 1986 as cited by Smart et al. 45 proposed an alternative typing system based on an immunodiffusion test that used antigens extracted from either the capsule or, in the case of unencapsulated strains, the outer membrane. They described seven serovars. Serovars 2 and 5 correspond to the serovars A and B respectively of Bakos et al.Kielstein and Rapp-Gabrielson in 1992 designated 15 serovars based on immunodiffusion using heat-stable antigens.17

Using either serotyping system, some strains cannot be typed.17, 35, 45 Smart et al. used restriction fragment length polymorphism (RFLP) analysis to group isolates of H. parasuis. 45 Thirteen groups were identified by this method. However, 24 of the 69 isolates examined did not correspond with any of the RFLP groups.

In a given herd, many strains of H. parasuis may be isolated, but in most cases, one or two strains predominate.45

Epidemiology

Infection with Haemophilus parasuis worldwide results in a sporadic disease affecting young pigs of 2 to 16 weeks of age. Single animals or litters may be affected, and the disease often involves the heaviest animals.29 When outbreaks occur they usually only involve a single farm, i.e. farm to farm spread is unusual. Specific pathogen-free (SPF) herds are more seriously affected by outbreaks and involve pigs of all ages.39 Morbidity rates are generally higher when older pigs are affected.31 In South Africa, the disease has been diagnosed more frequently in piglets two to six weeks of age, and often affects all the piglets in a single litter.10

Haemophilus parasuis commonly occurs among the flora of the upper respiratory tract and pharynx of healthy conventional pigs,12, 25, 38, 39, 43 but not of SPF pigs.39 Piglets acquire the organism soon after birth and immunity to Glässer’s disease has usually developed by the age of seven to eight weeks. The disease occurs only sporadically37, 43 apparently when certain conditions, particularly those associated with stress produced by transport, weaning, changes in diet or weather (especially chilling), movement to new quarters, and mixing of animals from different sources,29, 31 predispose animals to the disease. The disease is more common in winter than in summer.

Transmission is by direct contact, usually via the nasal route. 29 Experimentally, an acute disease with a short incubation period of less than 24 hours can readily be produced following intratracheal inoculation of H. parasuis. 24, 28 In addition, the condition can be reproduced by inoculation of the organism by other routes, such as intraperitoneal, intravenous and subdural.12, 38 Pathogenicity varies between strains.17, 20, 29 It has been found that when pigs were infected by intranasal inoculation and contact exposure under the same conditions, one strain induced severe fatal disease and another subclinical disease.2 The basis of virulence has not been established, but in one study it was demonstrated that virulence properties of strains are not directly correlated with epitopes based on lipo-oligosaccharides as antigens.52

Screening of herds by the bacteriological examination of nasal swabs will not necessarily indicate the probability of disease, as the strains isolated from nasal swabs are not always the same strains that cause disease in a herd.42

Pathogenesis

The pathogenesis of Glässer’s disease is incompletely understood.29 Piglets infected with H. parasuis soon after birth usually do not develop clinical disease because of the presence of maternal immunity. Under conditions of stress that may be due to environmental factors or co-infections, H. parasuis in vades the mucosa of the upper respiratory tract. Studies have indicated the nasal mucosa as the primary site of colonization, where microscopic lesions of acute suppurative rhinitis develop.50, 51 Focal loss of cilia and acute cell swelling, probably due to toxin release, were observed in the nasal and tracheal mucosa. Penetration of the surrounding tissues by H. parasuis results in haematogenous dissemination throughout the body. The bacterium has a predilection for serosal surfaces, such as those lining the large body cavities, joints and meninges, where it localizes to produce its pathological effects.37 The immune status of the host probably plays a paramount role in the establishment of the disease.29 This is supported not only by the age predilection for the disease, which is generally at the stage when maternally acquired antibodies of the piglet are waning, but also by the fact that SPF animals are more susceptible even in the absence of any apparent predisposing factors.31 Concurrent infection with certain viruses may exacerbate disease caused by H. parasuis. It has been found that piglets co-infected with porcine reproductive and respiratory syndrome (PRRS) virus and H. parasuis developed acute septicaemia rather than the more typical fibrinous polyserositis.47 Pigs concurrently infected with the virus that causes Aujeszky’s disease (pseudorabies) and a serovar of H. parasuis of medium virulence developed purulent pneumonia, possibly owing to destruction of respiratory epithelial cells by the virus that permitted the bacteria to proliferate in the lungs.27 Haemophilus parasuis can, however, act as a primary lung pathogen.19, 46

Studies have indicated that endotoxic shock and disseminated intravascular coagulation are important consequences of acute septicaemia induced by H. parasuis. 1, 3

Clinical signs

The duration of the incubation period of the disease under field conditions is not known, but it is likely to be short as clinical signs can become evident within two to three days of animals being stressed.11

The manifestations of the clinical disease depend upon the localization and severity of the lesions. In some animals, clinical signs may not be noticed before they simply expire suddenly.44 In those in which signs are apparent, the disease is generally of sudden onset, with a peracute to acute course of one to two days, and is characterized by fever with rectal temperatures as high as 42 °C. The affected animals also manifest anorexia, listlessness, respiratory distress (respiration is shallow and rapid, and may be accompanied by extension of the neck and opening of the mouth with breathing), increased heart rate, lameness and nervous signs.5, 12, 32, 34, 44, 48

Coughing may be present and auscultation of the thoracic cavity may reveal the presence of pleural and pericardial frictional rubbing sounds. The skin may develop cyanosis before death supervenes.

Most animals show some degree of lameness and develop a shuffling gait as the result of pain due to polyarthritis. One or several of the joints of the limbs, particularly the larger ones, are swollen, hot and tender. In an apparently atypical form of H. parasuis infection, sows became inappetent, weak and ataxic, and several developed head swelling, as a result of inflammation of the masseter muscles.14

Involvement of the nervous system is manifested in many animals by depression, stupor, hyperaesthesia, muscular tremors, ataxia, paresis or paralysis, and convulsions.29, 32

In piggeries where the disease occurs, the morbidity rate is variable29 but the mortality rate in untreated cases may be very high.12 Persistent disease in survivors may occur as chronic arthritis, meningitis or intestinal obstruction due to the presence of peritoneal adhesions.5, 48

Pathology

In typical cases of Glässer’s disease, the macroscopic pathology is characterized by the presence of a serofibrinous meningitis of the brain and spinal cord, pleuritis, pericarditis, peritonitis and polyarthritis,12 but not all of these lesions are equally evident or even present in all individuals; in some they are not present or may be difficult to detect, particularly in those that die of the peracute disease.38, 44

In the central nervous system, the meninges of the brain are generally more severely affected than are those of the spinal cord. The cerebrospinal fluid is frequently cloudy.12, 32

In animals suffering from polyarthritis, the atlantooccipital joint and the larger joints of the limbs are particularly involved.12, 32 In these joints the inflammatory reaction is characterized by an excessive amount of turbid synovial fluid, and the presence of yellowish-grey flecks or deposits of fibrinous exudate adhering to the synovial membranes or joint surfaces.32

The pleural, pericardial and peritoneal cavities contain small to larger amounts of serous fluid in which yellowish floccules of fibrin may be floating. The serosal surfaces may contain flecks and strands of fibrin that in some cases may be very delicate and difficult to detect unless one is aware of the possibility of their existence.12, 49 Mucosal congestion of the gastrointestinal tract, particularly in the fundus of the stomach, and multiple petechial haemorrhages in the renal cortices may be present. A few animals exhibit bronchopneumonia with or without the presence of the other typical lesions of Glässer’s disease.11 It has, however, been found that in endemic situations outbreaks in piglets and growers in large units were increasingly characterized by bronchopneumonia without serositis.16 These outbreaks were ascribed to high infection pressure and inadequate immunity. Acute purulent myositis involving the masseter muscles has been described in SPF sows.14

The histological changes reflect those of the gross pathology with, in addition, thrombosis of some vessels in the meninges, renal glomeruli and dermal papillae.32, 33, 38

Diagnosis

In a typical case of Glässer’s disease, the presence of serofibrinous polyserositis, polyarthritis and meningitis is sufficient to make a provisional diagnosis, which is often reinforced if the history includes the existence of a predisposing stress factor.

Affected tissues or swabs of the exudate should be submitted for bacterial isolation to confirm the diagnosis. The meningeal exudate and the cerebrospinal fluid seem to be the most suitable for this purpose.12 In most cases, the organism can be isolated from the lungs, but very seldom from other organs, such as the liver, kidneys and spleen.12 Swabs should be placed in a transport medium (such as Amies or Stuarts) for submission to a diagnostic laboratory as H. parasuis grows poorly and is susceptible to desiccation.

Generally, the isolation of H. parasuis is readily achieved12 unless the material from which cultures are made is grossly contaminated. Modified Levinthal agar (containing 1,6 units of bacitracin/ml), chocolate agar (containing 1 in 25 000 dilution of crystal violet and 1,6 units of bacitracin),23 or broth with V-factor added29 are suitable selective media.

Several researchers have investigated immunohistochemical techniques for the detection of H. parasuis and concluded that the techniques were useful, particularly in cases where culture was difficult, for example when the animals had been treated with antibiotics.1, 39 A range of tissues should be taken, including meninges, peritoneum, spleen and liver capsule, as cross-reactivity with Actinobacillus pleuropneumoniae in lung tissue may occur.39 For detection of H. parasuis in highly contaminated samples such as nasal swabs, an oligosaccharide-specific capture plate hybridization (OSCPH) test was found to be more sensitive and specific than culture.6

As most herds are subclinically infected, serological surveys usually indicate low complement fixation titres in most animals but high titres are suggestive of recent infection in fully susceptible pigs.30

Differential diagnosis

Mycoplasmal infections, such as that caused by Mycoplasma hyorhinis, may cause similar clinical signs and lesions but are usually milder and more chronic than Glässer’s disease.12, 32 Erysipelas may present a similar picture but the range of the ages of affected animals is usually wider. Streptococcus suis infections cause a similar disease picture to parasuis infection but the exudate is more purulent. Actinobacillus pleuropneumoniae infections may occasionally mimic Glässer’s disease.

Teschen and Aujeszky’s disease should also be considered in countries where these diseases occur,48 especially when nervous signs are present.

Control

Several agents may be used for the treatment or prophylaxis of Glässer’s disease, including penicillin, tetracycline, and potentiated sulphonamides such as trimethoprim.8 Tiamulin administered in drinking water proved successful in eliminating H. parasuis infection from pig herds in which strategic early weaning was practised.26 Treatment of clinical disease should be by parenteral administration of the agent, and it is imperative that it be commenced as early in the course of the disease as possible. In addition, all the animals in the group in which the disease has diagnosed should be treated whether or not clinical signs are apparent.29, 48 It has been suggested that, in practice, penicillin should be the treatment of choice, followed by tetracyclines.15 For chemoprophylaxis, the addition of one of the agents to feed or water can also be considered. It should be borne in mind, however, that strains with resistance to penicillin, tetracyclines and other chemotherapeutic agents have been encountered. For these reasons, the sensitivity of the organism should be continuously monitored in all herds which are experiencing disease as a result of H. parasuis infection.15, 16

Although vaccines are not generally available, they have been used to good effect.43 In one outbreak in a SPF herd in Denmark associated with a very high morbidity rate, prophylactic use of a formalized autologous vaccine prepared from cultures of H. parasuis and containing aluminium hydroxide as adjuvant, provided significant protection.31 A commercial vaccine (Glassinord R, Northern Drugs and Chemicals Ltd, Copenhagen) was subsequently developed and successfully used to control the disease in Denmark and Switzerland,37 but owing to poor cross-immunization between serotypes, immunoprophylaxis may not always be successful.18, 36 Vaccination of pregnant gilts or sows seems to offer the best protection to their piglets through colostrum-derived antibodies, and the need to administer a booster vaccination to piglets depends upon the affected age group in the herd.46 The use of bacterins prepared from homologous strains (autogenous bacterins) tends to give better results in this respect in herds in which outbreaks occur.42

Management techniques that separate piglets from carrier sows before loss of passive immunity, such as modified early weaning and strategic early weaning, have proved successful in eliminating H. parasuis infection.7

References

  1. AMANO, H., SHIBATA, M., KAJIO, N. & MOROZUMI, T., 1994. Pathologic observations of pigs intranasally inoculated with serovar 1, 4 and 5 of Haemophilus parasuis using immunoperoxidase method. Journal of Veterinary Medical Science, 56, 639–644.
  2. AMANO, H., SHIBATA, M., KAJIO, N. & MOROZUMI, T., 1996. Pathogenicity of Haemophilus parasuis serovars 4 and 5 in contact-exposed pigs. Journal of Veterinary Medical Science, 58, 559–561.
  3. AMANO, H., SHIBATA, M., TAKAHASHI, K. & SASAKI, Y., 1997. Effects on endotoxin pathogenicity in pigs with acute septicaemia of Haemophilus parasuis infection. Journal of Veterinary Medical Science, 59, 451–455.
  4. BIBERSTEIN, E.L. & WHITE, D.C., 1969. A proposal for the establishment of two new Haemophilus species. Journal of Medical Microbiology, 2, 75–78.
  5. BLOOD, D.C. & HENDERSON, J.A., 1963. (eds). Veterinary Medicine. 2nd edn. Baltimore: The Williams and Wilkins Company.
  6. CASAMIGLIA, M. & PIJOAN, C., 1998. An OSCPH technique for detection of Haemophilus parasuis in nasal swabs. Proceedings of the 15th IPVS Congress, Birmingham, England, 5–9 July 1998, 2, 143.
  7. CLARKE, L.K. 1998. New rearing technologies: Influence on health, growth and production economics of swine. Proceedings of the 15th IPVS Congress, Birmingham, England, 5–9 July 1998, 1, 281–288.
  8. EAVES, L.E., BLACKWELL, P.G. & FEGAN, M., 1989. Characterization and antimicrobial sensitivity of haemophil isolated from pigs. Australian Veterinary Journal, 66, 1–4.
  9. GLÄSSER, K., 1910. Untersuchungen über die Schweineseuche mit besonderer Berücksichtigung ihrer Atiologie und Patologie. Deutsche Tierärztliche Wochenschrift, 18, 729–733.
  10. HENTON, M.M., 1992. Unpublished data. Onderstepoort Veterinary Institute, Onderstepoort.
  11. HIRASAWA, H., SATHO, Y., MOROZUMI, K., MOROZUMI, T., NAKAGAWA, Y., OTA, T., TANAKA, K., OSADA, N. & TAKADA, T., 1987. Outbreak of Haemophilus parasuis infection in swine associated with purulent fibrinous meningitis. Journal of the Japan Veterinary Medical Association, 40, 519–522.
  12. HJARRE, A., 1958. Enzootic virus pneumonia and Glässer’s disease of swine. Advances in Veterinary Science, 4, 235–263. New York & London: Academic Press.
  13. HJARRE, A. & WARMBY, G., 1942. cited by Nielsen, R. & Danielsen, V
  14. HOEFLING, D.C., 1991. Acute myositis associated with Haemophilus parasuis in primary SPF sows. Journal of Veterinary Diagnostic Investigation, 3, 354–355.
  15. KIELSTEIN, P., 1985. Zur Glässerschen Krankheiten und Chemotherapeutika-Empfindlicheit ihres Erregers. Monatshefte für Veterinärmedizin, 40, 806–809.
  16. KIELSTEIN, P. & LEIRER, R., 1990. Zur Glässerschen Krankheit des Schweines — Ätiologisch-epidemiologische Untersuchungen zum Erregerspektrum. Monatshefte für Veterinärmedizin, 45, 577–582.
  17. KIELSTEIN, P. & RAPP-GABRIELSON, V.J. 1992. Designation of 15 serovars of Haemophilus parasuis on the basis of immunodiffusion using heat-stable antigen extracts. Journal of Clinical Microbiology, 30, 862–865.
  18. KIELSTEIN, P. & RASSBACH, A., 1991. Serologische Typisierung und Nachweiss immunoger Kreuzreaktionen von Haemophilus parasuis (Glässersche Krankheit). Monatshefte für Veterinärmedizin, 46, 586–589.
  19. KIELSTEIN, P., RASSBACH, A., POHLE, D., JOHANNSEN, U., WIEGAND, M. & SCHAFER, M., 1994. Zur Pathogenese der Haemophilus parasuis-Infektion des Schweines (Glässersche Krankheit). Monatshefte für Veterinärmedizin, 49, 71–75.
  20. KIELSTEIN, P., ROSNER, H. & RASSBACH, A., 1992. Die Glässersche Krankheit als Enzootie in Schweinegrossbeständen und die Bedeutung bestimmter biologischer Eigenschaften von Haemophilus parasuis für den Krankheitsverlauf. Monatshefte für Veterinärmedizin, 47, 539–544.
  21. KILIAN, M. & BIBERSTEIN, E.L., 1984. Genus II. Haemophilus. In: KRIEG, N.R. & HOLT, J.G., (eds). Bergey’s Manual of Systematic Bacteriology. Vol. 1. Baltimore: Williams & Wilkins.
  22. KRIEG, N.R. & HOLT, J.G., 1984. Bergey’s Manual of Systematic Bacteriology. Vol. 1. Baltimore: Williams & Wilkins
  23. LITTLE, T.W.A., 1970. Haemophilus infection in pigs. The Veterinary Record, 87, 399–402.
  24. LITTLE, T.W.A. & HARDIUNG, J.D.J., 1971. The comparative pathogenicity of two porcine Haemophilus species. The Veterinary Record, 88, 540–545.
  25. MOLLER, K. & KILLIAN, H., 1990. V factor-dependent members of the family Pasteurellaceae in the porcine upper respiratory tract. Journal of Clinical Microbiology, 28, 2711–2716.
  26. MOORE, C.A. & SANFORD, S.E. 1998. Control of Actinobacillus pleuropneumoniae pneumonia and possible eradication of Haemophilus parasuis from co-mingled, multi-sourced, SEW piglets after treatment with Tiamulin. Proceedings of the 15th IPVS Congress, Birmingham, England, 5–9 July 1998, 2, 166.
  27. NARITA, M., KAWASHIMA, K., MATSUURA, S., UCHIMURA, A. & MIURA, Y., 1994. Pneumonia in pigs infected with pseudorabies virus and Haemophilus parasuis serovar 4. Journal of Comparative Pathology, 110, 329–339
  28. NEIL, D.H., MCKAY, K.A., L’ECUYER, C.L. & CORNER, A.H., 1969. Glässer’s disease of swine produced by intratracheal inoculation of Haemophilus suis. Canadian Journal of Comparative Medicine, 33, 187–193.
  29. NICOLET, J., 1992. Haemophilus infections. In: LEMAN, A.D., STRAW, B., MENGELING, W.L., D’ALLAIRE, S. & TAYLOR, D.J., (eds). Diseases of Swine. 7th edn. Ames: Iowa State University Press.
  30. NIELSEN, R., 1981. Haemophilus infections in Danish swine herds. In: KILIAN, M., FREDERIKSEN, W. & BIBERSTEIN, E.L., (eds). Haemophilus, Pasteurella and Actinobocillus. London, New York, Toronto, Sydney, San Francisco: Academic Press.
  31. NIELSEN, R. & DANIELSEN, V., 1975. An outbreak of Glässer’s disease. Studies on the etiology, serology and the effect of vaccination. Nordisk Veterinaer Medicin, 27, 20–25.
  32. PALMER, N., 1993. Bones and joints. In: JUBB, K.V.F., KENNEDY, P.C., & PALMER, N., (eds). Pathology of Domestic Animals. 4th edn. Vol. 1. San Diego, New York, Boston, London, Sydney, Tokyo, Toronto: Academic Press
  33. PEET, R.L., FRY, J., LLOYD, J., HENDERSON, J., CURRAN, J. & MOIR, D., 1983. Haemophilus parasuis septicaemia in pigs. Australian Veterinary Journal, 60, 187.
  34. RADOSTITS, O.M., RUHNKE, H.L. & LOSOS, G.J., 1963. An outbreak of meningitis in swine caused by Haemophilus species or organism. Canadian Veterinary Journal, 4, 265–270.
  35. RAPP-GABRIELSON, V.J. & GABRIELSON, D. 1992. Prevalence of Haemophilus parasuis serovars among isolates from swine. American Journal of Veterinary Research, 53, 659–664.
  36. RAPP-GABRIELSON, V.J., KOCUR, G.J., CLARK, J.T. & MUIR, S.K., 1997. Haemophilus parasuis: Immunity in swine after vaccination. Veterinary Medicine, 92, 83–90.
  37. RIISING, H.J., 1981. Prevention of Glässer’s disease through immunity to Haemophilus parasuis. Zentralblatt für Veterinärmedizin, Reihe B, 28, 630–638.
  38. RILEY, M.G.I., 1977. Haemophilus parasuis infection in swine. Journal of the American Veterinary Medical Association, 171, 649–651.
  39. SÉGALES, J., DOMINGO, M., SOLANO, G.I. & PIJOAN, C., 1997. Immunohistochemical detection of H. parasuis serovar 5 in formalin-fixed paraffin-embedded tissues of experimentally infected swine. Journal of Veterinary Diagnostic Investigation, 9, 237–243
  40. SHANKS, P.L., 1939. Acute arthritis in pigs. The Veterinary Record, 51, 783–784.
  41. SHOPE, R.E., 1931. Swine influenza. I. Experimental transmission and etiology. Journal of Experimental Medicine, 54, 349–385
  42. SMART, N.L., HURNIK, D. & MACINNES, J.I., 1993. An investigation of enzootic Glässer’s disease in a specific-pathogen-free grower-finisher facility using restriction endonuclease analysis. Canadian Veterinary Journal, 34, 487–490.
  43. SMART, N.L. & MINIATS, O.P., 1989. Preliminary assessment of a Haemophilus parasuis bacterin for use in specific pathogen-free swine. Canadian Journal of Veterinary Research, 53, 390–393
  44. SMART, N.L., MINIATS, O.P. & FRIENDSHIP, R.M., 1989. Glässer’s disease and prevalence of subclinical infection with Haemophilus parasuis in swine in southern Ontario. Canadian Veterinary Journal, 53, 339–343.
  45. SMART, N.L., MINIATS, O.P. & MACINNES, J.I., 1988. Analysis of Haemophilus parasuis isolates from southern Ontario swine by restriction endonuclease fingerprinting. Canadian Journal of Veterinary Research, 52, 319–324.
  46. SOLANO-AGUILAR, G.I., PIJOAN, C., RAPP-GABRIELSON, V., COLLINS, J., CARVALHO, L.F. & WINKELMAN, N., 1999. Protective role of maternal antibodies against Haemophilus parasuis infection. American Journal of Veterinary Research, 60, 81–87.
  47. SOLANO, G.I., SEGALES, J., COLLINS, J.E., MOLITOR, T.W. & PIJOAN, C. 1997. Porcine reproductive and respiratory disease syndrome virus (PRRSv) interaction with Haemophilus parasuis. Veterinary Microbiology, 55, 247–257.
  48. TAYLOR, D.J., 1983. Pig Diseases. 3rd edn. Cambridge: The Burlington Press.
  49. TUSTIN, R.C., 1992. Faculty of Veterinary Science, University of Pretoria. Personal communication
  50. VAHLE, J.L., HAYNES, J.S. & ANDREWS, J.J., 1995. Experimental reproduction of Haemophilus parasuis infection in swine: Clinical, bacteriologic, and morphologic findings. Journal of Veterinary Diagnostic Investigation, 7, 476–480.
  51. VAHLE, J.L., HAYNES, J.S. & ANDREWS, J.J. 1997. Interaction of Haemophilus parasuis with nasal and tracheal mucosa following intranasal inoculation of cesarean derived colostrum deprived (CDCD) swine. Canadian Journal of Veterinary Research, 61, 200–206.
  52. ZUCKER, B.A., BAGHIAN, A., TRUAX, R., O’REILLY, K.L. & STORZ, J., 1996. Detection of strain-specific antigenic epitopes on the lipo-oligosaccharide of Haemophilus parasuis by use of monoclonal and polyclonal antibodies. American Journal of Veterinary Research, 57, 63–67.