2
Johne’s Disease in Domesticated and Wild Animals

Most information about Johne’s disease (JD) comes from dairy cattle. However, it is important to consider how the disease also is exhibited in other domesticated and wild animals. The significance of species other than dairy cattle as potential sources of exposure could increase as control programs are implemented and as within-herd transmission declines on dairy farms. Table 2– 1 summarizes the reported clinical indications of JD in domesticated and wild animal species. The following summary begins with dairy cattle, which reflect closely the condition in beef cattle, and concludes with the current state of knowledge about other species.

Table 2–1. Species Infected with Mycobacterium avium subsp. paratuberculosis

Species

Location

Reference

Domesticated ruminants

Cattle

Global

Buergelt et al., 1978b

Sheep

Global

Rajya and Singh, 1961

Goats

Global

Nakamatsu et al., 1968

Camels

Unspecified

Ganke et al., 1964



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2 Johne’s Disease in Domesticated and Wild Animals Most information about Johne’s disease (JD) comes from dairy cattle. However, it is important to consider how the disease also is exhibited in other domesticated and wild animals. The significance of species other than dairy cattle as potential sources of exposure could increase as control programs are implemented and as within-herd transmission declines on dairy farms. Table 2– 1 summarizes the reported clinical indications of JD in domesticated and wild animal species. The following summary begins with dairy cattle, which reflect closely the condition in beef cattle, and concludes with the current state of knowledge about other species. Table 2–1. Species Infected with Mycobacterium avium subsp. paratuberculosis Species Location Reference Domesticated ruminants Cattle Global Buergelt et al., 1978b Sheep Global Rajya and Singh, 1961 Goats Global Nakamatsu et al., 1968 Camels Unspecified Ganke et al., 1964

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Species Location Reference Reindeer Unspecified Katic, 1961 Llamas Unspecified Rankin, 1958 Zebu Cattle Unspecified Katic, 1961 Wildlife and exotic ruminants White-tailed deer (Odocoileus virginianus) Unspecified Chiodini and Van Kruiningen, 1983 Reindeer (Rangifer tarandus) Unspecified Katic, 1961 Sika deer (Cervus nippon) Unspecified Chiodini et al., 1984 Red deer (Cervus elaphus) Unspecified Hillermark, 1966 Axis deer (Axis axis) Unspecified Chiodini et al., 1984 Roe deer (Capreoleus capreoleus) Unspecified Katic, 1961 Fallow deer (Dama dama) Unspecified Chiodini et al., 1984 Moose (Alces alces) Unspecified Chiodini et al., 1984 Rocky Mountain bighorn sheep (Ovis canadensis) Colorado Quist, 1998; Williams et al., 1979 Aoudads (Ammotragus lervia) Unspecified Boever and Peters, 1974 Mouflons (Ovis musimon) Unspecified Boever and Peters, 1974 Rocky Mountain bighorn goats (Oreamnos americanus) Colorado Quist, 1998; Williams et al., 1979 Dwarf goats Unspecified Katic, 1960 American bison (Bison bison) Unspecified Chiodini et al., 1984 American buffalo (Syncerus caffer) Unspecified Katic, 1961 Water buffalo (Bubalus bubalis) Unspecified Katic, 1961 Bactrian camels (Camelus bactrianus) Unspecified Katic, 1961 Dromedary camels (Camelus dromedarius) Unspecified Amand, 1974 Antelopes Unspecified Katic, 1961 Stonebuck (Capra aegagraus ibex) Unspecified Williams and Spraker, 1979 Tule elk (Cervus elaphus nannodes) California Quist, 1998; Jessup et al., 1981 Llamas (Llama glama) Unspecified Appleby and Head, 1954 Yaks (Bos grunniens) Unspecified Almejan, 1958

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Species Location Reference Gnus (Connochaetes albojubatus) Unspecified Rankin, 1958b Zebu cattle (Bos indicus) Unspecified Katic, 1961 Wild rabbits Scotland Grieg et al., 1999; Beard et al., 2001b Monogastric animals Mules Global Eveleth and Gifford, 1943 Hogs Global Larsen et al., 1971 Chickens Global Larsen et al, 1981 Monkeys Global Chiodini et al., 1984 Pygmy asses Global Van Ulsen, 1970 Mandrills (Papio sphinx) Unspecified Zwick et al., 2002 Horses Global Larsen et al., 1972   SOURCE: Adapted from Chiodini et al., 1984. SPECTRUM OF DISEASE IN DOMESTICATED ANIMALS Cattle JD is characterized by vague and often variable clinical signs, and the clinical signs and the severity of gross and histological lesions do not always correspond (Allen et al., 1968; Downham, 1950; Hallman and Witter, 1933; Macindoe, 1950; Smyth, 1935; Smyth and Christie, 1950). Whitlock and Buergelt (1996) have described the following progression of disease in cattle (summarized in Table 2–2). Stage I: Silent Infection In Stage I, animals typically exhibit no overt evidence of infection with Mycobacterium avium subsp. paratuberculosis (Map). Stage I JD is typically found in calves and heifers, most immature young stock, and many adult cattle. No routine or special clinicopathologic tests or serology will detect disease in these animals. Only postmortum tissue culture or, less often, histopathology can detect infection at this early stage of disease. Stage II: Subclinical Disease Most animals in Stage II JD are adults that are carriers of Map. The animals do not exhibit clinical signs typical of JD, but they sometimes have detectable antibodies or exhibit altered cellular immune responses. Many are fecal-culture negative, although they intermittently shed low numbers of organisms in feces. In a small percentage (15–25 percent), disease can be detected by fecal culture, by altered cellular immune response, by serum antibodies, or by histopathology. An unknown proportion of Stage II animals progress slowly to Stage III (clinical disease), but because so many are culled

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from herds for other reasons and before clinical signs typical of JD are recognized, the magnitude of the Map infection within a herd can be obscured. Stage III:Clinical Disease The clinical signs characteristic of Stage III JD typically develop only after several years of Map incubation. The initial signs are subtle; they include a drop in milk production, roughening of the hair coat, and gradual weight loss despite an apparently normal appetite. Over a period of several weeks, diarrhea (often intermittent at first) develops. In the absence of a history of herd infection, clinical diagnosis is difficult because other conditions (gastrointestinal parasitism, peritonitis, renal amyloidosis, lymphosarcoma, copper deficiency, chronic salmonellosis, chronic infectious diseases, starvation, kidney failure) often result in similar signs. Because JD diagnosis based on the signs is challenging, the first cases in a herd often are misdiagnosed (Whittington and Sergeant, 2001). Histopathologic lesions can occasionally be found in the intestinal tract, with the most common site being the terminal ileum. Serum and plasma biochemical changes are predictable and characteristic of the clinical signs, but they are not specific enough to be of use in diagnosis of JD. Most animals test positive on fecal culture for Map and have detectable concentrations of antibodies on commercial serum enzyme-linked immunosorbent assay (ELISA) and agar gel immunodiffusion tests, although these diagnostic tests, discussed in Chapter 3, may misidentify animals. A few unusual cases will regress to Stage II and remain there for an indeterminate period. Stage IV:Advanced Clinical Disease Animals can progress from Stage III to Stage IV JD in a few weeks, and their health deteriorates rapidly. They become increasingly lethargic, weak, and emaciated as the disease progresses to Stage IV. Intermandibular edema (bottle jaw) due to hypoproteinemia, cachexia, and profuse diarrhea characterize Stage IV. Dissemination of Map throughout the tissues can occur. Although the organism can sometimes be cultured from sites distant from the gastrointestinal tract, extraintestinal lesions are rarely detected. When extraintestinal lesions are present, the liver, other parts of the GI tract, and the lymph nodes are the most common sites. At this stage, most animals are culled from the herd because of decreased milk production or severe weight loss. Those animals often are sent for salvage to slaughter, where their carcasses may not pass tests for human consumption. Death from JD is often the result of the severe dehydration and cachexia. At any given time in an infected herd, the majority of infected animals will be in Stages I and II, with relatively few animals exhibiting clinical signs of disease (Stages III and IV). Because the total number of animals with clinical signs grossly underestimates the prevalence of infection in the herd, the animals in Stages III and IV have been referred to as the “tip of the iceberg” (Whitlock and Buergelt, 1996).

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Table 2–2. Clinical Stages of Johne’s Disease in Cattle Stage Sign Presence of Histopathologic Lesion I Silent infection No overt evidence Absent or undetectable II Subclinical No clinical signs exhibited Sometimes III Clinical Production loss, roughening of hair coat, gradual weight loss despite normal appetite, diarrhea Can be found in intestinal tract; most common site: terminal ileum IV Advanced clinical Lethargy, weakness, emaciation, intermandibular edema due to hypoproteinemia, cachexia, profuse diarrhea Can be found in organs other than intestinal tract; most common secondary site: liver and lymph nodes Sheep and Goats Clinical disease in sheep and goats is reported to be similar to that in cattle, with the exception that diarrhea is less frequent and onset occurs in younger animals. When diarrhea does occur, it typically attends end-stage disease (Smith and Sherman, 1994; Stehman, 1996; Williams et al., 1983). In a study of 50 clinically affected sheep—45 of which were emaciated and 5 of which were in fair condition—50 percent had hard feces; 30 percent had soft, non-pelleted feces; and 20 percent had diarrhea (Carrigan and Seaman, 1990). Wool fragility, gaps in wool, weak fibers (metabolic alopecia subsequent to malnutrition or severe disease), and poor fleece condition have also been reported in sheep with JD (Cranwell, 1993). Diseases with similar clinical signs in small ruminants include chronic intestinal parasitism, internal abscesses such as those caused by Corynebacterium pseudotuberculosis, chronic hepatic disease, and chronic malnutrition (Beeman et al., 1989). Clinical signs thus are not a reliable indicator of the presence or absence of Map infection in sheep or goats (Whittington and Sergeant, 2001). Moreover, Map until recently was difficult to isolate from feces of infected sheep (Juste et al., 1991); advances in culture technique, however, appear to have largely solved that problem (Whittington et al., 1999). SPECTRUM OF DISEASE IN OTHER ANIMALS Clinical manifestations of JD in non-domesticated ruminants are highly variable in prevalence and timing, but otherwise are similar to those of their domesticated counterparts (Manning, 1998; Power et al., 1993; Ridge, 1991; Williams et al., 1979, 1983). Little has been published specifically on the clinical signs of JD in non-ruminant wildlife, but Map-infected European rabbits rarely exhibit signs (Beard et al., 2001b; Greig, 1997).

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PATHOLOGY Cattle Lesions of Subclinical Infection Gross lesions in Stage I and Stage II JD cattle are usually absent or undetectable. The earliest histologic lesions observed in experimentally infected calves are focal aggregates of epithelioid macrophages or Langerhans giant cells in Peyer’s patches and in the villous tips of the intestine (Buergelt et al., 1978b; Gilmour et al., 1965; Payne and Rankin, 1961a, b). Similar histologic changes and involvement of mesenteric lymph nodes also characterize naturally infected animals in subclinical stages of disease. Acid-fast staining of lesions may reveal a few mycobacteria, but staining often is negative. Lesions of Clinical Infection Primary gross lesions in cattle are confined to the intestinal tract, where they can extend from the duodenum to the rectum. The terminal ileum is the most common site. Gross lesions can be subtle, but the affected intestine is usually thickened, corrugated, and highly folded. In animals infected with strains of Map that produce pigment, the intestinal mucosa can take on a yellow-orange cast (Stuart, 1965a, b; Taylor, 1951, 1953; Watt, 1954). The regional lymph nodes also can be involved, sometimes with enlarged and tortuous mesenteric lymphatics. Common secondary changes include effusion into body cavities, atrophy of fat, general wasting, and dependent subcutaneous edema. The histologic lesion generally is described as a diffuse granulomatous or histiocytic enteritis, without necrosis, hyperemia, or reactionary fibrosis (Buergelt et al., 1978b; Hallman and Witter, 1933; Taylor, 1953). Early granulomatous infiltrates in the intestine are often nodular (tuberculoid), and, in later stages of disease, they can coalesce to form diffuse (lepromatous) infiltrates (Chiodini et al., 1984). The villous tips of the mucosa often fuse, reducing the surface area and the absorptive capacity of the intestine (Whitlock and Buergelt, 1996). The lamina propria and submucosa of the intestine are variably infiltrated with macrophages. Giant cells, primarily of the Langerhans type, often are observed. Epitheloid macrophages and giant cells can contain variable numbers of typical acid-fast bacilli (Buergelt et al., 1978b; Hallman and Witter, 1933; Harding, 1957; Nguyen and Buergelt, 1983; Rajya and Singh, 1961; Stamp and Watt, 1954). Affected lymph nodes exhibit a granulomatous lymphadenitis. Granulomatous lymphangitis also can occur, although less so in cattle than in sheep and goats. In the terminal stages of disease, lesions can occur in other organs; the liver is the most common secondary site (Buergelt et al., 1978b; Hallman and Witter, 1933; Mathews, 1930; Whitlock and Buergelt, 1996).

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Sheep Several descriptions of the lesions of JD in sheep have been published (Carrigan and Seaman, 1990; Clarke, 1997; Clarke and Little, 1996; Perez et al., 1996; Rajya and Singh, 1961; Stamp and Watt, 1954). The pathology of naturally occurring and experimental cases in sheep has been extensively reviewed by Clarke (1997). Gross lesions are generally similar to those described for cattle, with thickening and corrugation of intestinal mucosa, thickening of lymphatic vessels, and lymph node enlargement. Lesions are more prevalent in the terminal ileum, but they often extend to other parts of the small intestine and colon. Carcass emaciation, intermandibular edema, ascites, hydropericardium, and serous atrophy of fat are common secondary findings. Several ovine strains of Map produce large amounts of an orange-yellow pigment that can result in brown-yellow pigmented intestinal lesions (Clarke, 1997). Lesion classification systems have been developed for JD in sheep. One of the more comprehensive systems designed to represent stages in the pathogenesis of the disease was described by Perez and colleagues (1996) and reviewed by Clarke (1997). The following summary of this system is adapted from Clarke (1997) and Perez and colleagues (1996). Category 1 This category has mild focal aggregates of foamy macrophages, forming small granulomas in the ileal Peyer’s patches with no visible organisms or visible gross lesions. The researchers indicated that this stage is exhibited in animals that were infected as lambs but in which disease has been arrested by effective cell-mediated immunity. Categories 2 and 3 represent disease progression into adulthood, with failure of effective immune mechanisms. Category 2 Animals in this category exhibit more extensive lesions in the ileal Peyer’s patches, with granulomas extending into the submucosa, and with obvious presence of organisms histologically but no visible gross lesions. Category 3a. Grossly visible thickening of the intestinal mucosa is present in this category. Multifocal large granulomas are in the lamina propria, submucosa, and serosa of the ileum and in draining lymph nodes, with extension of lesions into the jejunum. Organisms are obvious histologically. Category 3b. In this category numerous macrophages and a few multinucleate giant cells spread in mosaic-like sheets through the submucosa and lamina propria to create villous fusion and marked thickening of the intestine with abundant organisms. Category 3c. A diffuse granulomatous enteritis is present in this category, with marked lymphocytic infiltrates within the mucosa and small, well-defined granulomas and giant cells scattered throughout the lesions.

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Organisms are either sparse or undetectable. Granulomas and focal areas of giant-cell necrosis are present within mesenteric lymph nodes. Goats Various breeds of domesticated and exotic goats are susceptible to Map infection. Subclinical paratuberculosis has been described in goats after experimental infection (Storset et al., 2001). Gross lesions in goats can be variable in type and location (Clarke, 1997; Stehman, 1996). The most common gross lesions are thickening of the terminal small intestine, mesenteric lymph node enlargement, and corrugation of the ileal mucosa. The earliest histologic lesions in experimental cases consisted of clusters of epithelioid macrophages and giant cells in the basal regions of ileal and jejunal Peyer’s patches at three months after inoculation. Lesions tended to coalesce and had extended into the large intestine by 10 months, when mucosal ulceration without caseous necrosis was evident (Clarke, 1997). In some cases, nodular foci of caseous necrosis with mineralization have been described in the mucosa, submucosa, serosa, lymphatics, and particularly in the mesenteric lymph nodes, which could be easily confused with signs typical of M. bovis or M. tuberculosis infection (Clarke, 1997). There is some evidence that subspecies other than Map, such as M. avium subsp. silvaticum, could be the etiologic agent in some caprine cases, possibly accounting for some of the prominent nodular caseonecrotic lesions that resemble tuberculosis (Collins et al., 1984; Thorel et al., 1990a). In advanced cases, the lesions of granulomatous enteritis are similar to those seen in other ruminants, but goats also exhibit lesions in the sciatic and brachial plexus nerves that resemble those in human leprosy. Granulomatous lesions also are present in the liver and lungs of some goats with advanced clinical disease (Stehman, 1996). A classification system for the lesions of JD in goats has been proposed (Corpa et al., 2000) that closely follows the system described above for sheep (Perez et al., 1996). Exotic Ruminants JD in small ruminants, deer, and South American camelids (llamas, alpacas) has been reviewed by Stehman (1996), Clarke (1997), and Collins et al. (1994). As in cattle, transmission of Map in sheep, goats, South American camelids, and deer is presumed to occur primarily through the fecal-oral route. Gross and microscopic lesions in South American camelids are similar to those reported in cattle, but lymph node necrosis and mineralization, along with multiorgan dissemination, have also been reported (Stehman, 1996). Gross and microscopic lesions in deer are similar to those reported in sheep and goats (Stehman, 1996). M. bovis and M. avium subsp. avium infections, which produce gross and histologic lesions indistinguishable from JD, have been reported in deer (Stehman, 1996).

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Non-ruminant Species Primates Map infection produced lesions confined to the intestine and abdominal lymph nodes in a colony of stump-tailed macaques at Yerkes Primate Center, resulting in a condition very similar to JD in cattle and Crohn’s disease in humans (McClure et al., 1987). Thirty-eight animals in the colony were infected and were shedding Map in feces. Using a serum ELISA, antibodies to Map could be detected in 79–84 percent of the animals in the colony; antibodies could not be detected in six animals with clinical disease (McClure et al., 1987). Map also was recently reported in a mandrill (Papio sphinx) (Zwick et al., 2002). Rabbits European rabbits (Oryctolagus cuniculi) infected with Map develop intestinal disease that is similar to ruminant JD (Angus, 1990). Laboratory rabbits, which are derived from the European rabbit, are occasionally used as an experimental model (Mokresh et al., 1989). Severe lesions in rabbits consist of extensive granulomas and numerous giant cells, with many intracellular acid-fast bacilli in the small intestine (Beard et al., 2001a). Other Species Monogastric species—including pigs, horses, dogs, and other laboratory animals—have demonstrated susceptibility to infection with Map under natural or experimental conditions (Chiodini et al., 1984; Clarke, 1997; Hines et al., 1995; Thoen et al., 1977, 1981), although reports of such cases are rare. Lesions in these species generally consisted of granulomatous enteritis and mesenteric lymphadenitis with the presence of organisms, but often without clinical signs. Occasional disseminated disease also occurs (Clarke, 1997). EPIDEMIOLOGY Global Prevalence in Domesticated Animals No systematic global survey has been completed for JD or for the presence of Map in domesticated animals, but JD has been reported on every continent of the world except Antarctica ([International Office of Epizootics] OIE, 2001b). Some countries, principally island nations, have reported no cases of JD, and others have reported JD limited to specific geographic zones. This self-reporting, however, requires no documentation and has not been independently verified. OIE, which consists of the veterinary administrations of 158 countries, has provided international standards and guidance for the JD diagnostic techniques, vaccines, and biologics (OIE, 2000). However, determining worldwide prevalence with any degree of certainty is complicated by the lack of international consensus on population-testing protocols.

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Furthermore, no international scientific consensus exists concerning the definition and verification of JD-free zones or regions (Table 2–3). Numerous JD prevalence studies have been completed in domesticated-animal populations of various countries (Kennedy and Benedictus, 2001). Differences in sampling design and diagnostic strategies make direct comparison of the studies difficult, but critical evaluation of published surveys suggests considerable variation in the prevalence of infected herds in different countries and within specific geographic areas. The prevalence of Map-infected animals and clinical JD also varies greatly within affected herds. Several estimates for JD herd prevalence exist (Tables 2–3 and 2–4). The most extensive surveys for JD and Map across multiple states in the United States have been completed by the United States Department of Agriculture (USDA). Merkal and colleagues (1987) cultured ileocecal lymph node specimens from clinically normal cull cattle at slaughterhouses in 32 states during 1983–1984. They isolated Map from 119 of 7540 cattle; isolates from dairy cattle culls were more than 250 percent higher than were those from beef culls (2.9 percent and 0.8 percent, respectively). USDA’s National Animal Health Monitoring System (NAHMS) serologically tested a sample of dairy and beef cattle using ELISA to obtain estimates of the prevalence of JD in herds and in individual animals. In the U.S., 32,622 cows were tested from 1004 dairy operations. Herd prevalence for JD was estimated at 21.6 percent (NAHMS, 1997a). Testing of 10,371 beef cows from 380 herds in 1997 generated an estimate of 7.9 percent infected herds, although the results are not directly comparable to those from dairy cattle because of differences in the investigators’ definitions of a positive test in a herd (Dargatz et al., 2001). Nevertheless, the surveys confirm the beliefs that JD is widespread in the United States and that dairy herds are more likely to be infected than are beef herds. Prevalence data for sheep in the United States will be available when the NAHMS Sheep 2001 report is released. No prevalence studies have been conducted on goats in the United States. Table 2–3. Global Seroprevalence of Johne’s Disease in Dairy Cattle Country Herd Prevalence (%) Australia (Victoria) 14–17 New Zealand 60 Netherlands 55 Belgium 22 Austria 7 England, Wales 17 Germany (Arnesberg) 10–30   SOURCE: Collins and Manning, 2002b.

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Table 2–4. Seroprevalence of Mine’s Disease in U.S. Dairy Cattle in 1996 Region Herd Prevalence (%) Westa 23.5 Midwestb 24.2 Northeastc 16.1 Southeastd 17.2 aOregon, Washington, Idaho, California, Texas, New Mexico. bMinnesota, Iowa, Missouri, Wisconsin, Illinois, Indiana, Ohio, Michigan. cVermont, New York, Pennsylvania. dFlorida, Tennessee, Kentucky. SOURCE: Adapted from Garry et al., 1999. Prevalence in Wildlife and Zoo Animals Although Map is primarily known as a pathogen of cattle and other domesticated ruminants, natural and experimental Map infection of ruminant and non-ruminant wildlife has been well documented (Angus, 1990; Beard et al., 2001a, b; Buergelt et al., 2000; Chiodini and van Kruiningen, 1983; Cook et al., 1997; Dukes et al., 1992; Hillermark, 1966; Jessup et al., 1981; Libke and Walton, 1975; McClure et al., 1987; Nebbia et al., 2000; Riemann et al., 1979; Shulaw et al., 1986; Stehman, 1996; Williams et al., 1983). However, the prevalence of JD in wildlife in the United States has not been investigated. Quist (1998) summarized the reports of JD in wildlife in the United States and identified only two endemic foci: Rocky Mountain bighorn sheep (Ovis canadensis) and mountain goats (Oreamnos americanus) in Colorado (Williams et al., 1979), and tule elk (Cervus elaphus nannodes) in California (Cook et al., 1997; Jessup et al., 1981). This could be interpreted as evidence of low JD prevalence in U.S. wildlife, but the lack of active surveillance suggests that data are insufficient to warrant any conclusions. Sporadic cases or outbreaks of JD have been reported in exotic hoofstock in U.S. zoos (Boever and Peters, 1974; Dukes et al., 1992; Weber et al., 1992) and game farms (de Lisle et al., 1993; Fawcett et al., 1995; Manning et al., 1998; Power et al., 1993). However, the prevalence of JD in zoos could be higher than is suggested by the paucity of reports. A recent survey of zoos accredited by the American Zoo and Aquarium Association revealed that 31 percent of responding institutions had experienced at least one case of JD, but up to 67 percent lacked effective surveillance for JD (Manning and Ziccardi, 2000). This suggests that true institutional prevalence is much higher. No prevalence studies have been conducted on game-ranched or farmed non-domesticated ruminants.

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plausible, as it is general knowledge that Map requires an acid medium for effective growth in culture. Pasteurization With concerns focused on transmission within and across species (especially cattle to humans), reports of Map in raw and processed milk (Grant et al., 1996, 1999, 2002a, 2002b; Millar et al., 1996; Streeter et al., 1995; Sweeney et al., 1992b; Taylor et al., 1981) have raised questions about the capability of heat treatment, pasteurization, and other processes to eliminate live organisms in colostrum and milk. Several researchers have examined this method of breaking the potential cycle of transmission through milk and dairy products (Table 2–6). Data indicate that Map is remarkably tolerant of a wide range of environmental conditions. Consistent with these data, early studies indicated that Map were more thermally resistant than were other milkborne pathogens (Chiodini and Hermon-Taylor, 1993; Sung and Collins, 1998). At relatively low inoculation concentrations and at a range of temperature and time treatments, live Map organisms were still detectable after incubation (Chiodini and Hermon-Taylor, 1993; Grant et al., 1996; Meylan et al., 1996; Rowe et al., 2000) (Table 2–6). They are known to form colonial clumps, and the clumped colonies demonstrate higher thermal resistance than do declumped samples, which has been attributed to the protective function of the outer layer of clumped cells allowing the inner layers to survive for longer heating times (Rowe et al., 2000). However, Map generally is more susceptible to even low-temperature treatments, when those treatments extend over long periods (5–30 min vs. 15 s) (Stabel, 2001). Map also can be inactivated by ionizing irradiation (Stabel et al., 2001). Should Map be identified as a significant zoonotic pathogen, pasteurization of milk will be a potential control action that can address a major point of human exposure. Studies currently leave the question of the effectiveness of heat treatment unresolved because of methodologic differences and the difficulty of quantifying viable Map organisms, and further work is warranted (Collins, 1997). A more pressing question is whether to implement changes in pasteurization and sterilization procedures before the zoonotic potential of Map has been clarified. This question was recently addressed in deliberations of the United Kingdom Food Standards Agency. Their report reached the conclusion that a link between Map and Crohn’s disease can neither be proved nor disproved with available evidence. However, they felt it important to take the possibility of such a link seriously. They included a recommendation that minimum pasteurization times be extended from 15 to 25 minutes. It was acknowledged that the evidence supporting this recommendation remains incomplete.

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Table 2–6.Mycobacterium avium subsp. paratuberculosis Response to Heat Treatment, Pasteurization, and Other Processes Inoculation Treatment Treatment Type Duration Incubation Period Result Reference Waste milk n/a 150°F Heat 30 min 28 weeks Complete inactivation Stabel, 2001 Milk 104 cfu/mL 0 kGy Ionizing radiation 1.2 kGy/h Unspecified Complete inactivation Stabel et al., 2001 104 cfu/mL 5 kGy Ionizing radiation 1.2 kGy/h Unspecified Complete inactivation Stabel et al., 2001 104 cfu/mL 10 kGy Ionizing radiation 1.2 kGy/h Unspecified Complete inactivation Stabel et al., 2001 108 cfu/mL 0 kGy Ionizing radiation 1.2 kGy/h Unspecified Complete inactivation Stabel et al., 2001 108 cfu/mL 5 kGy Ionizing radiation 1.2 kGy/h Unspecified Complete inactivation Stabel et al., 2001 108 cfu/mL 10 kGy Ionizing radiation 1.2 kGy/h Unspecified Complete inactivation Stabel et al., 2001 Colostrum 104 cfu/mL 63°C Heat 30 min 16 weeks Reduction Meylan et al., 1996 104 cfu/mL 20–23°C n/a Unspecified 16 weeks Reduction Meylan et al., 1996 103 cfu/mL 63°C Heat 30 min 16 weeks Reduction Meylan et al., 1996 103 cfu/mL 20–23°C n/a Unspecified 16 weeks Reduction Meylan et al., 1996 102 cfu/mL 63°C Heat 30 min 16 weeks Reduction Meylan et al., 1996 102 cfu/mL 20–23°C n/a Unspecified 16 weeks Reduction Meylan et al., 1996 Raw cow’s milk 106 cfu/mL 72°C Heat 15 sec Unspecified Reduction Grant et al., 1999 106 cfu/mL 75°C Heat 15 sec Unspecified Reduction Grant et al., 1999 106 cfu/mL 78°C Heat 15 sec Unspecified Reduction Grant et al., 1999

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Inoculation Treatment Treatment Type Duration Incubation Period Result Reference 106 cfu/mL 80°C Heat 15 sec Unspecified Reduction Grant et al., 1999 106 cfu/mL 85°C Heat 15 sec Unspecified Reduction Grant et al., 1999 106 cfu/mL 90°C Heat 15 sec Unspecified Reduction Grant et al., 1999 106 cfu/mL 72°C Heat 20 sec Unspecified Incomplete inactivation Grant et al., 1999 106 cfu/mL 72°C Heat 25 sec Unspecified Complete inactivation Grant et al., 1999 107 cfu/mL 63.5°C Heat 30 min Unspecified Reduction Grant et al., 1996 102 cfu/mL Unspecified Unspecified Unspecified Unspecified Reduction Grant et al., 1996 104 cfu/mL 71.7°C Heat 15 sec Unspecified 3–100% Inactivation Chiodini and Hermon-Taylor, 1993 >103 cfu/mL 71.7°C Heat 15 sec Unspecified Reduction Grant et al., 1996 Raw cow’s milk, w/clumpedMap Unspecified 63°C Heat 0 min 18 weeks at 37°C Reduction Rowe et al., 2000 Unspecified 63°C Heat 2 min 18 weeks at 37°C Reduction Rowe et al., 2000 Unspecified 63°C Heat 4 min 18 weeks at 37°C Reduction Rowe et al., 2000 Unspecified 63°C Heat 6 min 18 weeks at 37°C Reduction Rowe et al., 2000 Unspecified 63°C Heat 8 min 18 weeks at 37°C Reduction Rowe et al., 2000 Unspecified 63°C Heat 10 min 18 weeks at 37°C Complete inactivation Rowe et al., 2000 Raw cow’s milk, w/declumpedMap Unspecified 63°C Heat 0 min 18 weeks at 37°C Reduction Rowe et al., 2000 Unspecified 63°C Heat 1 min 18 weeks at 37°C Reduction Rowe et al., 2000 Unspecified 63°C Heat 2 min 18 weeks at 37°C Reduction Rowe et al., 2000

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Inoculation Treatment Treatment Type Duration Incubation Period Result Reference Unspecified 63°C Heat 3 min 18 weeks at 37°C Reduction Rowe et al., 2000 Unspecified 63°C Heat 4 min 18 weeks at 37°C Reduction Rowe et al., 2000 Unspecified 63°C Heat 5 min 18 weeks at 37°C Complete inactivation Rowe et al., 2000 Notes: cfu/mL, colony forming units/per milliliter; kGy, kiloGray. PATHOGENESIS IN CATTLE The severity of disease and the rate of its progression depend on several variables, among the most important of which are the dose of Map and the age of the animal at exposure. Only a small dose of the organism might be required to establish infection in a newborn calf, even though large doses might be necessary to infect older animals (reviewed in Sweeney, 1996). The incubation period for JD also is variable but often protracted, ranging from 4 months to 15 years (Macindoe, 1950; Smyth and Christie, 1950). In oral transmission, after ingestion, the organism is taken up by specialized M cells lining the intestine (Gilmour et al., 1965; Momotani et al., 1988; Payne and Rankin, 1961a, b). The mechanism by which attachment and uptake of Map occurs is not known, but recent in-vitro studies suggest that soluble fibronectin binds to fibronectin attachment proteins on both Map and epithelial cells to mediate uptake (Secott et al., 2001, 2002). After uptake by the M cells, Map bacilli are transferred to underlying lymphoid tissue. Dissemination via the bloodstream can then occur, with subsequent localization to secondary sites—the liver, spleen, and peripheral lymph nodes. The organism is ultimately taken up by macrophages, where it survives and is isolated from normal humoral and cellular defense mechanisms (Momotani et al., 1988; Zurbrick and Czuprynski, 1987). The mechanism by which Map survives within macrophages has not been elucidated, but it has been shown with other mycobacteria that the normal phagosome-lysosome fusion with acidification and the release of hydrolytic enzymes does not occur (Armstrong and Hart, 1971, 1975; de Chastellier and Thilo, 1999; Frehel et al., 1986a; Goren et al., 1987a, b; Hackam et al., 1998; Hart and Young, 1988; Pieters, 2001a; Sibley and Krahenbuhl, 1987; Sibley et al., 1987). The mechanism by which this inhibition occurs is unclear, but recent work suggests that two host cell components, the steroid cholesterol and a phagosomal coat protein called TACO (tryptophane aspartate-containing), promote the establishment of an intracellular infection by mycobacteria (Ferrari et al., 1999; Pieters, 2001a). Cheville and colleagues (2001) suggested that pathogenic and wild strains of Map block phagosomal acidification, so that the phagosome fails to obtain markers of the late phagosome and phagolysosome. This then leads to

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replication within bacteriophorous vacuoles (Cheville et al., 2001). Their study and a previous study by Draper and Rees (1973) show that, during growth within phagosomes of murine or bovine macrophages, the mycobacteria develop a capsule-like lamellar coat. This coat has an ultrastructural organization that is compatible with mycoside (a mycobacterial glycopeptidolipid), which could be an intraphagolysosomal defense mechanism (Cheville et al., 2001; Frehel et al., 1986b; Rastogi et al., 1994; Rulong et al., 1991). Iron is a requirement for mycobacterial growth. However, unlike other mycobacteria, Map does not produce siderophores (mycobactins) that facilitate iron uptake. In-vitro growth of Map therefore requires supplementation with mycobactin. The mechanism by which Map organisms obtain iron in vivo is not known, but recent studies suggest that Map produces its own ferric reductase enzyme that mobilizes host iron to allow growth (Homuth et al., 1998). In animals that fail to contain the infection, replication within tissues progresses, and more and more of the intestinal lining (mucosa) and regional lymph nodes are infiltrated by macrophages filled with mycobacteria. This phase of infection corresponds with the prolonged incubation period observed clinically. Eventually, the accumulating mycobacteria-laden macrophages interfere with intestinal absorption, resulting in weight loss and an initially intermittent diarrhea. It has been proposed that formation of mycobacterial antigen-antibody complexes in the infected intestine results in histamine release, thus exacerbating the diarrhea (Merkal et al., 1970). Failure of the immune system to contain the infection results in a continuously increasing mycobacterial burden and progressively more severe clinical disease. In the terminal stages of the infection, immune cells become functionally nonresponsive, resulting in uncontrolled replication and spread of Map in tissues (Chiodini et al., 1984). The mechanism by which this unresponsiveness occurs in the advanced stages of the disease is unknown and has not been thoroughly investigated. The release of soluble mycobacterial products, bacterial-cell components, or mediators from macrophages could be involved, as in M. leprae infections (Birdi et al., 1980; Salgame et al., 1980; Zurbrick et al., 1988). Age-Related Susceptibility Reports from experimental inoculations in cattle suggest that infection must occur early because of the development of age-related resistance (Bendixen, 1978; Doyle, 1953, 1956; Hagan, 1938; Larsen et al., 1975; Levi, 1948; Rankin, 1958a, 1961a, b, 1962; Taylor, 1953). The mechanism by which age-related resistance occurs is still not known. It has been proposed that the greater susceptibility in neonates is related to increased permeability of the intestine in the first 24 hours after birth, when colostral immunoglobulins are being absorbed. Alternatively, an inherent weakness of the intestinal mucosal barrier at this time could be involved (Sweeney, 1996). One study suggests that by one year of age, resistance appears equal to that of mature adult cattle (Buergelt et al., 1978b). Recent work has shown that there is a differential distribution of γδ-T-lymphocyte subpopulations between young and mature cattle (Wyatt et al., 1994). It has been suggested that cytotoxic inactivation of

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antigen specific CD4+ helper lymphocyte populations by γδ-T-cells could be involved in the proliferation of Map and in the development of clinical disease (Chiodini and Davis, 1993). Some of the mechanisms underlying age-related resistance have been explained, but there are still questions about the importance of horizontal transmission in older animals. Age-related susceptibility to Map infection has not been investigated thoroughly in species other than cattle. Genetic Susceptibility Genetic susceptibility to JD has been investigated only in mice, cattle, and humans. The susceptibility of mice to the establishment of mycobacterial infections has been shown to be controlled by a single, dominant, autosomal gene called the Bcg gene locus, which directly regulates the process of T-cell-dependent macrophage activation for antimycobacterial function and, indirectly, the quality and magnitude of the specific immune response to M. bovis (Skamene, 1989). It is thought that the Bcg-resistant allele confers to the macrophage the ability to inhibit the proliferation of mycobacteria (Frelier et al., 1990). Similar genetic resistance has been described for M. intracellulare (Goto et al., 1984). Inbred C57BL/6J mice are more susceptible to infection by Map than are outbred Swiss mice, and resistance to Map and M. bovis is regulated by the same locus or by linked loci (Frelier et al., 1990). The Nramp gene (a component of the Bcg locus) has been linked to resistance in mycobacterial infections, including murine models of JD (Blackwell et al., 1994; Chandler, 1961; Frelier et al., 1990; Hackam et al., 1998; Levin and Newport, 2000; Veazey et al., 1995). The bovine equivalent of that gene has been identified, and based on its homology to the murine Nramp1 gene, it could have similar functions (Feng et al., 1996). However, Barthel and colleagues (2000) were unable to detect an association between resistance or susceptibility to infection with M. bovis and polymorphism in the Nramp1 gene, suggesting that the Nramp1 gene might not determine resistance or susceptibility to M. bovis infections in cattle. The involvement of the Nramp1 gene in the elimination of Map in cattle is currently unknown (Valentin-Weigand and Goethe, 1999). Mutations in other genes critical for macrophage up-regulation, including interferon-γ (IFN-γ), IFN-γ receptor, tumor necrosis factor (TNF) receptor, or interleukin-12 (IL-12), or IL-12-receptor genes, are all associated with decreased resistance to mycobacterial infection in humans (Levin and Newport, 2000). Studies of genetic resistance could be important in the control of JD if they promote the identification or development of resistant cattle (Koets et al., 2000). Immune Response Efforts to dissect the immune system and identify the mechanisms that regulate the immune response to pathogens have involved the use of multiple laboratory species, especially the mouse. The use of inbred mouse strains with inherited differences in susceptibility to infectious agents, and knockout mice missing genes encoding molecules involved in development of an immune

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response, have greatly advanced our knowledge of the composition of the immune system and mechanisms of immune regulation. The use of other species has provided further information on factors regulating immune responses, and also revealed there are species differences in the composition of the immune system that need to be taken into consideration when attempting to understand the immune response to a pathogen in the target species (Davis and Hamilton, 1998). This is especially important for studies in ruminants (Davis et al., 1996; Goddeeris, 1998; MacHugh et al., 1993; Wijngaard et al., 1994). Three major lineages of lymphocytes have been identified: αβ- and γδ-T-lymphocytes, and B lymphocytes. The subset composition of the αβ-T and B lymphocyte lineages is similar in most species (Goddeeris, 1998). However, the composition of the γδ-T-lymphocyte lineage differs. In most species, there is one lineage of γδ-T-lymphocytes that is present in low frequency (three to five percent) in peripheral blood, but widely distributed in mucosal tissue at sites of entry of pathogens. There are two lineages of γδ-T-lymphocytes in ruminants and other Artiodactyla (pigs and camelids) that differ in phenotype and tissue distribution (Davis et al., 1996, 1998, 2000; Goddeeris, 1998; MacHugh et al., 1998). One population with a phenotype similar to γδ-T-lymphocytes in humans and mice is present in blood (three to five percent) and tissues in comparable proportions, except in the spleen where they may comprise 30 percent or more of the lymphocytes present (Davis et al., 1996). The second population is distinguished by the expression of a unique molecule, workshop cluster 1 (WC1). It differs from the WC1− population in frequency in peripheral blood and in the pattern of trafficking, potentially associated with differences in function (Wilson et al., 1998, 1999). The WC1+ population may comprise 30 to 50 percent of lymphocytes in peripheral blood of young animals. Except for the spleen, the WC1+ and WC1− γδ-T-lymphocytes are present in similar proportions in secondary lymphoid tissue and in epithelial tissues at points of entry of pathogens (Wyatt et al., 1994, 1996). Additional smaller subsets of lymphocytes have also been identified, including natural killer cells (NK). Limited information is available on the role of these subsets and γδ-T-lymphocytes in host defense (Kaufmann, 1996). However, it is thought that γδ-T-lymphocytes and NK cells may play a role in first line of defense against infectious agents (Kaufmann, 1996). The development of a protective immune response is complex (Seder and Hill, 2000; Van Parijs and Abbas, 1998). It involves the interaction of multiple cell types following exposure to a pathogen. In general, there are four phases to the response: (1) antigen recognition following encounter with a pathogen, (2) increase in the frequency of antigen-specific lymphocytes involved in cell-mediated immune (CMI) response and humoral immunity, (3) contraction of the responding populations through apoptosis following control of the infection, and (4) appearance of memory lymphocytes. Protective immunity is dependent on the development and maintenance of memory cells following encounter with pathogens. Immunity is lost if the concentration of antigen-specific memory cells drops below a threshold level needed for a rapid recall

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response on subsequent encounters with a pathogen (Seder and Hill, 2000; Sprent and Surh, 2001). Cumulative studies have shown αβ-T- and B-lymphocytes are the primary mediators of immunity. Their function is under the control of dendritic cells (DC), the sentinel antigen presenting cells (APC) in peripheral lymphoid tissue that capture and process pathogen derived antigens for presentation to CD4+ and CD8+ subsets of αβ-T-lymphocytes (Banchereau and Steiman, 1998; Flores-Romo, 2001; Huang et al., 2000; Iwasaki and Kelsall, 2000; Kelsall and Strober, 1996). Following uptake of antigen at sites of infection, DC migrate to secondary lymphoid organs, where they present signatory peptide fragments of antigens to CD4+ and CD8+ T-lymphocytes in association with major histocompatibility complex (MHC) class II or I molecules, respectively. Second signals mediated through co-stimulatory molecules expressed on DC, and secreted chemical messengers (chemokines and cytokines) modulate differentiation and maturation of T-lymphocytes to cells with specific effector and/or memory activity (Banchereau and Steinman, 1998; Dubois et al., 1998; Flores-Romo, 2001; Moser and Murphy, 2000). In particular, secretion of IL-12 by DC promotes differentiation of CD4+ lymphocytes with a type 1 cytokine profile dominated by secretion of IFN-γ (Bloom, 1992; Dubois et al., 1998). Lymphocytes with this cytokine profile develop into effector cells that function in CMI and provide help in activation of B cells that produce complement-fixing antibodies. IFN-γ secreted by type 1 CD4+ lymphocytes activates macrophages and increases their capacity to kill ingested bacteria. Direct killing of bacteria is mediated by secretion of perforin and granulysin at the sites of infection (Canaday et al., 2001; Dieli et al., 2001; Smyth et al., 2001). Down-regulation of secretion of IL-12 promotes differentiation of CD4+ lymphocytes with a type 2 cytokine profile dominated by secretion of IL-4. Lymphocytes with this cytokine profile function as helper cells that stimulate activation of B cells that produce antibodies without complement fixing activity. DC also promote differentiation of CD4+ lymphocytes with a type 3 cytokine profile dominated by expression of IL-10 and TGF-β. Lymphocytes with this cytokine profile modulate the maturation of lymphocytes with type 1 and type 2 cytokine profiles by down regulating their effector activity. Stimulation of CD8+ T-lymphocytes by DC promotes differentiation of cytotoxic T-lymphocytes (CTL) with a type 1 cytokine profile and/or T-lymphocytes with a type 2 cytokine profile that regulate CD4+ and CD8+ T-lymphocyte effector activity. The role of DC in the response of γδ-T-lymphocytes is not clear, but may involve the interaction with another MHC related molecule, CD1 (Spada et al., 2000). A subset of γδ-T-lymphocytes in humans has been shown to be selectively activated through CD1c. This subset contains perforin and granulysin and, when activated, secrete IFN-γ and IL-2 (Spada et al., 2000). Granulysin-dependent killing has also been reported for γδ-T-lymphocytes expressing a specific arrangement of the γδ-T-cell receptor (Dieli et al., 2001). The magnitude and type of immune response depends on the pathogen and in part on which receptors are used for internalization of the pathogen and which signaling pathways are used for antigen processing (Huang et al., 2001;

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Lutz et al., 1996; Pulendran et al., 2001; Thoma-Uszynski et al., 2000, 2001). Some pathogens have evolved ways to persist in APC and modulate the pathways of antigen processing and secretion of cytokines for a sufficient time to cause disease (Gewurz et al., 2001; Pieters, 2001a). In spite of these various strategies to evade immune elimination, either or both cellular and humoral responses develop against pathogens. The response elicited may lead to either resolution and sterile immunity, or to immune control of infection, with the pathogen persisting in a dormant state or replicating at a low rate. Elucidation of how different pathogens influence the evolution of an immune response is crucial to understanding how to develop efficacious vaccines against known and emerging pathogens (Zinkernagel, 2000). The most informative data on the immune response to mycobacteria have been obtained from studies that used M. tuberculosis, M. bovis and M. leprae in humans, and M. tuberculosis and M. bovis in mice and cattle. The limited Map studies have shown the factors regulating the immune response could be similar to those that regulate other mycobacteria. Common features include tropism for macrophages and use of the same receptors for internalization and sequestration in a phagosomal compartment that does not fuse with lysosomes. Once the mycobacteria take up residence in macrophages, several immunologic mechanisms become involved in the control or containment of the infection. Studies with humans and mice with genetic defects in genes encoding IFN-γ, TNF-α, IL-1, IL-12, and their receptors have shown the mechanisms of those cytokines and receptors in the development and maintenance of an immune response that controls infection (Dorman and Holland, 2000; Jouanguy et al, 1999; Ottenhoff, 2000). Genetically susceptible mice and knockout mice with defects in the αβ and γδ-T-cell receptors, CD4, and CD8 have shown that CD4+ memory T-lymphocytes with a type 1 cytokine profile (Bloom, 1992) operate in the control of infection (Follett and Czuprynski, 1990; Hamilton et al., 1991; Koets et al., 2002; Mogues et al., 2001). The effector mechanisms used by CD4+ T-lymphocytes include secretion of IFN-γ, which activates bactericidal activity in macrophages; secretion of lymphotoxin, which promotes formation and maintenance of tuberculoid granulomas at sites of infection (Roach et al., 2001); and secretion of perforin and granulysin, which kill bacteria on contact (Canaday et al., 2001). The role of CD8+ memory and γδ-T-lymphocytes in protective immunity is less clear but there is evidence that CD8+ T-lymphocytes also kill M. tuberculosis using granulysin (Canaday et al., 2001). Their presence in lesions suggests they synergize with CD4+ T-lymphocytes. Some studies suggest that the type-1 (Bloom, 1992) cell-mediated immune response described above contains the infection during the early, subclinical stages (Demangel and Britton, 2000; Koets et al., 2002). The type-1 immune response can be recognized histologically by the presence of microscopic granulomas and infiltrating lymphocytes with very few mycobacteria, and it is known as the tuberculoid form of the disease. The mechanisms by which mycobacteria ultimately evade host responses and replicate in macrophages are still being elucidated, but one important factor appears to be retention of the TACO protein in the macrophage phagosome,

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which in some way protects replicating bacteria from bactericidal oxygen and nitrous oxide intermediates produced by activated macrophages (Pieters, 2001b; Raupach and Kaufmann, 2001). Progression to clinical disease is thought to occur when the type-1 response declines, allowing proliferation and dissemination of mycobacteria. This stage of the immune response is characterized histologically by diffuse lesions (rather than discrete granulomas) and large numbers of Map, and it is called the lepromatous form of the disease (Burrells et al., 1999; Perez, 1996, 1999; Stabel, 2000). Additional work is needed to elucidate the factors that contribute to the decline in type-1 immune response and progression of disease, but a constellation of host factors and microbial virulence factors is likely involved. No vaccine has been developed that elicits an immune response that completely eliminates viable Map from the host (sterile immunity). This is in part attributable to a lack of knowledge of the factors that regulate the immune response to Map or to pathogenic mycobacteria in general (Flynn and Chan, 2001). What is known is that not all animals exposed to Map progress to clinical disease. It is not clear whether infection is never established in some animals, or whether they develop an immune response that controls or eliminates the pathogen. The appearance of a strong CMI response and the secretion of IFN-γ by cultures of peripheral blood lymphocytes stimulated with Map antigens may prove to be the earliest indications that animals have been permanently or transiently infected with Map (Köhler et al., 2001; Gwozdz and Thompson, 2002). If infection has occurred and is under immune control, CMI and secretion of IFN-γ by activated T-lymphocytes could be the only indicators of latent, persistent infection (Koets et al., 2002; Köhler et al., 2001). It is not possible, however to use those indicators to distinguish between animals that have developed sterile immunity and those with latent infections. If immune control is lost in latently infected animals, disease can progress slowly through subclinical to clinical stages, with associated lesions. The rate of progression would depend on the factors that influence the immune response to Map antigens. The indicators of disease progression are a depression in the CMI response to Map antigens and the appearance of a strong humoral antibody response to Map antigens (Chiodini, 1996; Koets et al., 2002; Stabel, 1998). Commercial and experimental live-attenuated and killed vaccines have elicited a CMI response that has not prevented infection or shedding of bacteria (Harris and Barletta, 2001; Kalis et al., 2001). Experimental trials with vaccinated animals challenged with virulent Map have, however, shown that immunization does have an effect. Vaccination leads to a reduction in the severity of lesions and bacterial load and to a reduction in clinical signs of disease (Harris and Barletta, 2001). The fact that some protective immunity is elicited suggests that further research could reveal which bacterial antigens to pursue for development of an efficacious vaccine. Sequencing of theMap Genome Complete sequencing of the Map genome is now at an advanced stage. This joint USDA National Animal Disease Center and University of Minnesota

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project has benefited from parallel efforts to sequence the genome of M. avium. Preliminary reports from the Map sequencing project demonstrate overall 99 percent sequence identity between Map and MAC. The Map genome size, at 5.5 million base-pairs, appears larger than that of M. tuberculosis and M. avium. Twenty-one unique genes had been identified as of January 2002 and it is projected that a total of about 50 will be identified when the project is complete. Among these is a new gene cluster without homology to any known genes at both the genome and possibly at the protein level. Preliminary expression studies have been started comparing expressed genes from culture vs. organisms cultivated in an immortalized macrophage cell line. Several repetitive elements have also been identified in addition to IS900 and IS1311. As might be expected, there are variations in sequence length between different copies of IS900 suggesting heterogeneity in this insertion element. This may be of value in more refined studies of strain relatedness. Results of the gene-sequencing project have the promise to provide diagnostic reagents with improved sensitivity and specificity. Expression libraries may help to identify unique antigens that can be used in developing serological assays and tests of cell-mediated immunity. They may also provide the foundation for rational vaccine development. These diagnostic approaches may additionally provide adequate specificity and sensitivity to assess definitively the role of Map in human disease.