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Incidence of Mycobacterium Avium Subspecies paratuberculosis in Mehsani and Surti Goats of Indian Origin using Multiple Diagnostic Tests

D.B. Barad, B.S. Chandel, A.I. Dadawala, H.C. Chauhan, H.S. Kher, S. Shroff, A.G. Bhagat, S.V. Singh, P.K. Singh, A.V. Singh, J.S. Sohal, S. Gupta, K.K. Chaubey, S. Chakraborty, R. Tiwari, R. Deb and K. Dhama

Mycobacterium avium subspecies paratuberculosis (MAP) is the causative agent of chronic enteric granulomatous inflammation in animals and is known as Johne’s Disease (JD) or Paratuberculosis. JD, being spectral in nature, presents variable bacteriological, immunological and pathological spectra leading to variable efficacy of diagnostic methods at different points of time during the course of infection. The present study aimed to estimate the incidence of MAP in two important breeds of goats (Mehsani and Surti) from South Gujarat region of India by applying conventional, molecular and serological methods. A total 219 goats were screened and categorized into Group-I (123 Mehsani goats), Group-II (76 Surti goats) and Group-III, (20 Non-descript goats). Percent positivity by faecal smear examination, delayed type hypersensitivity (DTH), agar gel immunodiffusion (AGID), IS900 polymerase chain reaction) (PCR) and indigenous enzyme linked immunosorbent assay (ELISA) kit was 9.2 (7/76), 21.9 (27/123), 10.9 (24/219), 12.5 (5/40) and 43.3% (95/219), respectively. Of the 123 goats of Group-I, 27 (21.9%) were positive in DTH test. Of the 5 faecal positive goats which also showed clinical signs, 2 (3.5%) goats died during study were negative by Johnin test. Similar to these findings, sensitivity of Johnin test in goats ranged between 18-30% with least specificity in both preclinical and advanced stage of disease. Of 34 cases of caprine paratuberculosis, 73.5% goats were positive for Johnin test. In the present study, out of the 5 infected goats, 3 (60%) were positive in Johnin test. Rectal pinch smear examination was carried out in 27 DTH positive goats and all smears were negative for the presence of acid fast bacilli. Screening tests (Indigenous ELISA and DTH) showed very high incidence of MAP infection in the goat population. The utility of multiple diagnostic tests is suggested for confirmatory detection and epidemiological diseases investigations of MAP in animals.

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D.B. Barad, B.S. Chandel, A.I. Dadawala, H.C. Chauhan, H.S. Kher, S. Shroff, A.G. Bhagat, S.V. Singh, P.K. Singh, A.V. Singh, J.S. Sohal, S. Gupta, K.K. Chaubey, S. Chakraborty, R. Tiwari, R. Deb and K. Dhama, 2014. Incidence of Mycobacterium Avium Subspecies paratuberculosis in Mehsani and Surti Goats of Indian Origin using Multiple Diagnostic Tests. Journal of Biological Sciences, 14: 124-133.

DOI: 10.3923/jbs.2014.124.133

Received: October 25, 2013; Accepted: November 11, 2013; Published: January 11, 2014


Mycobacterium avium subspecies paratuberculosis (MAP) is the causative agent of chronic enteric granulomatous inflammation in animals known as Johne’s disease (JD) or Paratuberculosis (Chiodini et al., 1984; Tripathi et al., 2002; Sweeney et al., 2012). It is either a saprophytic or obligatory microbe which is an ancient pathogen (Deb and Goswami, 2011). It causes entero-colitis, lymphadenitis and lymphangitis in domestic and wild ruminants (Perez et al., 1996; Clarke, 1997; Collins et al., 2001; Singh et al., 2012). The disease has an endemic distribution globally reducing production of food in both developed as well as developing countries. The disease in most of the instances is restricted to sub-clinical infection. But there is rapid loss in condition leading to quick death because of nutritional deficiency resulting due to absorption loss from the gut (Collins, 2003). In India, limited studies (Pande, 1940; Singh et al., 1996, 2007a) have reported the variable incidence (2-18%) in domestic livestock (Tripathi et al., 2002). Animals may either receive infection from infected parents through semen, in-utero, milk or colostrums (Shankar et al., 2010) or pick up infection from contaminated environment soon after birth. Then it follows a protracted incubation period which may last few months to years. In this century, MAP is considered as a major pathogen of animals as well as human beings (Manning and Collins, 2001; Tripathi et al., 2002; Momotani, 2012; Sweeney et al., 2012). The disease is primarily being responsible for low per animal productivity in the ruminant population (domestic) of the country. In the domestic livestock inspite of high prevalence and endemicity of the MAP the impact of the disease has neither been realized nor estimated at the National level. This is primarily due to the lack of indigenous diagnostic tests as well as kits. Sharing of MAP between species (inter-species transmission) has been frequently reported (Singh et al., 2012). Clinical symptom being non-specific (weight loss and diarrhea) are not visible before the disease gets fully established in the body (clinical). Progressive weakness ultimately leads to either un-timely death or culling. Internally, granulomatous enteritis leads to thickening of intestinal mucosa and regional lymphangitis and lymphadenitis in small ruminants (Chiodini et al., 1984). However, corrugations are rare as animals goes out much before from the production system. Recently, antibiotic resistance is an also an emerging problem to be tackled for effective treatment of the disease (Tiwari et al., 2013a).

Johne’s disease (JD) being spectral in nature presents variable bacteriological, immunological and pathological spectra leading to variable efficacy of diagnostic methods at different points of time during the course of infection (Chiodini et al., 1984). Cultivation of MAP from faeces and tissues (ileo-caecal lymph nodes and small intestines) though specific but is time consuming and may require up to 12 weeks (Cocito et al., 1994) and presents lesser sensitivity (Sockett et al., 1992). Polymerase Chain Reaction (PCR) is a rapid diagnostic test for the detection of MAP from faeces, blood and tissues with sensitivity comparable with bacterial culture (Tripathi et al., 2002; Singh et al., 2010; Dhama et al., 2011). Monoclonal antibody based diagnostics have also been reported to be useful for JD (Deb et al., 2013). Infected animals elicit strong Cell-Mediated Immune (CMI) response in the early stages of infection and strong humoral immune response in the later stages (Clarke, 1997; Perez et al., 1999). CMI based tests such as cutaneous testing with johnin Purified Protein Derivative (PPD), gamma-interferon assay and lymphocyte stimulation test, although reported to be sensitive, have been less frequently evaluated for the detection of early stages of MAP infection in sheep and goats (Molina et al., 1991; Storset et al., 2001; Kurade et al., 2004). Antibody assays for JD include Complement Fixation Test (CFT), Agar Gel Immunodiffusion (AGID) and Enzyme-linked Immunosorbent Assay (ELISA) tests (Kurade, 1999). The latter is sensitive in clinical infection but performs poorly in sub-clinical stages of infection (Stewart et al., 2006). In view of these facts, this study aimed to evaluate the efficacy of commonly used diagnostic tests including of conventional, serological and recent molecular tools for the detection of MAP infection in goats.


Faecal smear: About 2-3 g of faecal sample were collected from each of the goats directly from rectum, brought to the laboratory on the same day and processed. One gram of faecal sample was properly crushed with the help of sterilized pestle and mortar in sterilized distilled water. Mixture was then centrifuged at 2,600 rpm for 45 min. After centrifugation, supernatant was discarded and from the semisolid middle layer smear was prepared on a clean slide. Smear was air dried, heat fixed and stained with Ziehl-Nielsen (ZN) method and examined under microscope. Positive faecal samples were stored at -20°C for DNA extraction and culture.

Rectal pinch smear: Rectal pinch smear were examined in goats positive in Johnin test. Rectal pinch was collected with a sterilized artificial insemination sheath and smears were prepared. Smears were air dried, heat fixed, stained with Zeihl Nielson (ZN) method and observed under microscope. Smears exhibiting presence of short acid fast bacilli were considered positive for MAP.

Delayed type hypersensitivity (DTH)/johnin test: Single Intra-dermal (DTH) test was performed on 123 goats of group-I by inoculation of 100 μg of Johnin PPD, obtained from Division of Biological Products, Indian Veterinary Research Institute (IVRI), Izatnagar (India) in the form of Heat Concentrated Synthetic Medium (HCSM) of Mycobacterium paratuberculosis. Test was carried out as per the instructions of manufacturer on the side of neck. Skin thickness was measured with Vernier Calipers at pre, 24, 48 and 72 h post inoculation. Animals showing hot, edematous, painful skin thickness of more than 3 mm after 48 h were considered positive.

Post mortem and histopathology: Goats died of natural infection were subjected to post mortem examination. Tissues (Ileo-caecal junction of intestine, ileum and mesenteric lymph nodes) exhibiting gross lesions were collected and stored at -20°C without adding any preservative for DNA isolation and PCR. For histopathology, tissues were stored in 10% formalin at room temperature. Formalin fixed tissues were cut into thin (2-3 mm) pieces and washed thoroughly with water for several hours before putting in ascending grades of alcohol for dehydration. Dehydrated tissues were cleared in xylene, embedded in paraffin blocks and 5 micron thick sections were prepared (Luna, 1968). Sections were stained with haematoxylin and eosin (H and E) and Ziehl Nielsen method. Smears made from tissues (Ileocaecal junction, mesenteric lymph nodes) were heat fixed and stained with ZN method for the presence of acid fast bacilli indistinguishable to MAP.

Agar gel immunodiffusion (AGID) test: The protocol of AGID was followed as per Ferreira et al. (2002). Agarose was dissolved at 0.75 in 0.85% NaCl solution and buffered to pH 9.0 with 0.01 M tris (hydroxymethyl)-aminomethane and sodium azide 0.02%. Gel was poured in plates on a 4 mm thick layer and wells of 5 mm diameter and in a hexagonal pattern of six peripheral wells for serum samples and a central well for antigen were made. Protoplasmic Antigen (PPA) (Allied Monitor, USA) was used as antigen at the concentration of 10 mg mL-1. Positive serum was placed into wells adjacent to test sera. Diameter between the centers of all the adjacent wells was 8 mm. After the wells were filled plates were incubated in covered, moistened glass dishes to avoid drying. Plates were examined after 24 and 48 h of incubation at room temperature. Appearance of one or more clearly definable precipitation lines before or at 48 h constituted a positive test result. Absence of any precipitation lines was recorded as a negative test result. Serum from clinically infected animals was used as positive control.


From tissues: After collection of tissues (mesenteric lymph nodes and intestines) these were grounded and treated with 0.9% HPC (Hexa decylpyridinium chloride) overnight. The sediment (0.5-1.0 mL) was taken in 2.0 mL capacity eppendorf tubes and washed 3-4 times with Phosphate Buffered Saline (PBS) by spinning and vortexing. Pellet was subjected to DNA isolation as per Van Embden et al. (1993) method with some modification. Briefly, washed pellet was suspended in 1.0 mL 1X Tris EDTA (TE) buffer (pH 8.0) and centrifuged at 8,000 rpm for 15 min. Pellet was re-suspended in 450 μL 1X TE buffer and subjected to freezing and thawing (heating to boiling and snap cooling at -20°C) and the process repeated 3-4 times. Then, 40 μL of lysozyme was added and the tubes incubated at 37°C for 2 h. Proteinase K (6 μL) and 10% Sodium Dodecyl Sulphate (SDS) (56 μL) were added and the tube incubated at 65°C for 30 min. After that 64 μL of CTAB+80 μL 5M NaCl were added and incubated at 65°C for 30 min. Equal volume of Chloroform+Isoamyl alcohol (24:1) was added, mixed properly and centrifuged at 10,000 rpm for 10 min. DNA was precipitated by chilled absolute ethanol and washed with 70% chilled ethanol. DNA pellet was re-suspended in 30 μL 1X TE buffer and stored at -20°C until further use.

From faeces: Faecal samples were centrifuged at 2,600 rpm for 45 min and bacilli were concentrated in the interface of solid and watery layers. After discarding the top layer, semi-solid middle layer was collected by sterilized swab and stirred in tubes containing 40 mL 0.9% HPC for decontamination (18-24 h) at room temperature. For DNA isolation, 0.5-1.0 mL of sediment, after discarding the decontaminated layer was kept into a 2.0 mL capacity sterilized eppendorf tubes. Sediment was washed with PBS, 3-4 times by spinning and vortexing. Pellet was subjected to DNA isolation as per method described by Van Embden et al. (1993) with some modification.

Polymerase chain reaction (PCR): DNA was isolated from decontaminated fecal and tissue samples were subjected to specific IS900 PCR as per Green et al. (1989) and Vary et al. (1990). Briefly, PCR was set in volume of 25 μL, using 1.0-5.0 ng template DNA, 12.5 μL of 2X PCR Master Mix (Genei, Banglore) and 1.0 μL of each primer (10 pmole). Total of 35 cycles were performed in a thermocycler (MJ Research) for complete amplication reaction (Table 1). Thermal cycling conditions for primer 1 were: Initial denaturation at 94°C for 3 min, followed by 35 cycles of denaturation at 94°C for 10 sec, annealing at 50°C for 10 sec, extension at 72°C for 10 sec and final extension at 72°C for 3 min. Thermal cycling conditions for primer 2 were: Initial denaturation at 94°C for 4 min, followed by 35 cycles of denaturation at 94°C for 30 sec, annealing at 64°C for 30 sec, extension at 72°C for 30 sec and final extension at 72°C for 5 min (Table 2, 3 and 4). Presence and yield of specifc PCR product was analyzed by 1.5% agarose gel electrophoresis.

Table 1: Details of primers used for PCR

Table 2: Faecal and rectal pinch smear examination
D: Goats died during study, *Two goats found negative by PCR, n: No. of animals

Table 3: Delayed type hypersensitivity (DTH) /Johnin test
n: No. of animals

Table 4: Enzyme linked immuno sorbent assay
n: No.of animals

ELISA testing: Serum samples were screened by ‘indigenous ELISA kit’ as per the method of Singh et al. (2007b). Briefly, 10 μg plate-1 of MAP ‘Indian Bison type’ protoplasmic antigen was taken in 10 mL of carbonate-bicarbonate buffer (pH 9.6) and coated in duplicate wells of flat bottom 96 well ELISA plates. Plates were incubated at 4°C overnight, washed thrice with washing buffer (PBS+0.05% Tween-20, pH 7.4- PBST), blocked (3% skimmed milk powder in PBS) and incubated at 37°C for 1 h. Plates were washed thrice with PBST and stored at 4°C till further use. M. phlei (Allied Monitor Inc., USA) was mixed in serum dilution buffer before the dilution of each test serum sample and plate used for serum dilution was incubated for one hr at 37°C. After incubation, 100 μL of each adsorbed test serum was transferred to duplicate wells of ELISA plate and plates incubated for 2 h at 37°C. Then the plates were washed thrice with PBST. After washing, 100 μL of optimally diluted (1:8000) rabbit anti-goat horse radish peroxidase conjugate was added to wells and incubated at 37°C for 1 h. Plates were washed thrice with PBST and 200 μL of freshly prepared substrate Orthophenyl diamine (OPD), 5 mg plate-1 in substrate buffer (pH-5.0) was added to each well and incubated in dark for 30 min at room temperature. Absorbance was read at 450 nm in an ELISA reader.

Data analysis and statistics: Data were managed in Tables created using Microsoft Office Excel; Microsoft Corp., Redmond, Washington, USA. Positive and strong positive samples were considered positive for MAP infection in ELISA as per Collins (2002) by converting the OD values into sample-to-positive (S/P) ratio (using formula; Sample OD-Negative OD/Positive OD-Negative OD in Excel cells). Relative sensitivity, specificity, kappa values and proportional agreement were calculated comparing all the five tests with each other (Table 1 and 2).


In animals and human beings, infection of Mycobacterium avium subspecies paratuberculosis have been associated with chronic inflammation of the intestines and mesenteric lymph nodes in animals (JD) and Inflammatory Bowel Disease (IBD) or Crohn’s disease in human beings (Sweeney et al., 1998; Sechi et al., 2001, 2005).

Table 5: Agar gel immuno diffusion (AGID)
n: No.of animals

Table 6: IS 900 Polymerase chain reaction (PCR)
n: no.of animals

For the diagnosis of MAP infection, the choice of the diagnostic test is essential for the success of the control programs but it depends on the purpose and should invariably involve multiple tests in case of chronic MAP infections (Singh et al., 2009a). In the faecal smear examination, out of 56 goats of Group-I, 5 (8.9%) goats were positive, whereas, in Group-III, out of 20 goats, 2 (10.0%) goats were positive. Of a total of 76 goats screened, 7 (9.2%) were positive (Table 5) for MAP. Out of 5 faecal positive goats of Group-I, 3 died during the study. Clinical signs were observed in all the 7 faecal smear positive goats and these were also positive in ELISA and AGID. Tripathi et al. (2006) showed that out of 36 known cases of caprine paratuberculosis diagnosed by clinical and faecal smear examination, 72.2% goats were shedding MAP bacilli. Faecal smear examination with acid-fast staining detected more goats as positive as compared to bacterial culture and PCR (Munjal et al., 2007). Screening of 71 animals (55 goats and 16 sheep) belonging to Central Institute for Research on Goat (CIRG) located at Makhdoom (India) revealed that 40% goats and 31.2% sheep were shedding MAP in faeces (Singh et al., 2007a, b).

Rectal pinch smear examination was carried out in 27 DTH/Johnin positive goats. All the rectal pinch smears were negative for the presence of MAP. In accordance to this, OIE (2000) has described that the smears of rectal mucosa biopsies are challenging to get and may not be very helpful because the disease process is less likely to involve the rectum in sheep and goats ( Of the 123 goats of Group-I, 27 (21.9%) were positive in DTH test. Out of 5 faecal positive goats which also showed clinical signs, 2 (3.5%) goats died during the study were negative by Johnin test. Similar to these findings, Paliwal and Rajya (1982) and Rajukumar (1998) stated that sensitivity of Johnin test in goats ranged between 18-30% with least specificity in both preclinical and advanced stage of disease.

Fig. 1: Mehsani goat breed from south Gujarat

Tripathi et al. (2006) reported that out of 34 cases of caprine paratuberculosis, 73.5% goats were positive for Johnin test. In the present study, of 5 infected goats, 3 (60%) were positive in Johnin test. For screening of MAP antibodies, ELISA is a better choice as it improves the rate of detection or better estimation of MAP infection. There is moreover significant increase in the rate of detection (Braun et al., 1990). Of the 219 goats tested, 95 (43.3%) were positive by ELISA. Of 123 Mehsani goat breed (Group-I) (Fig. 1), positive reactors were higher 57 (46.3%) and had history of recurrent diarrhea (Table 6). Sweeney et al. (1995) reported that sensitivity of ELISA was only 15.0% in animals excreting low quantities of MAP bacilli in their faeces as compared to 87.0% in animals with clinical signs of Johne’s disease. In this study, similar results were obtained i.e., all the 7 goats showing clinical signs as well as faecal smear positivity were also positive by ELISA. Paolicchi et al. (2003) and Singh et al. (2009b) observed that absorbed ELISA was useful to detect positive animals and those goats shedding MAP in faeces. Similarly, in the present study all the animals shedding MAP in feces were positive in the ELISA test also. Indigenous ELISA kit developed by CIRG, Makhdoom was most sensitive and specific test for the detection of MAP infection (OIE, 2000).

As far as the AGID test is concerned it is well proven that animals either in the early stage or advanced stage of the disease are often found negative and there is every possible of chance of obtaining either false positive or false negative result (Stabel, 1998). Of 219 goats tested in the present study, 24 (10.9%) were positive by AGID. Group-III consisting of 20 non-descript goats showed higher number (4 or 20%) of goats as positive reactors. All the 7 goats showed clinical signs and were positive in smear examination and AGID. Similarly, Sherman et al. (1984) recorded that out of 33 AGID positive cattle, 32 (96.9%) were confirmed for MAP infection by culture or necropsy. Similarly, Ferreira et al. (2002) found AGID specificity was 92.5% and sensitivity was 57%. Mainar-Jaime et al. (1998) recorded 11.7% (64 of 546) sero-prevalence of MAP in sheep and goat flocks using AGID test.

PCR was carried out on faecal samples of suspected animals of Group-I and III with history of recurrent diarrhoea or pasty faeces and tissue samples of three goats died in Group-I. Of 40 goats, 7 showed clinical signs and were also positive by faecal smear examination (Fig. 2). Among t hese 7 goats, faecal PCR detected 5 goats as positive while 2 were negative. Munjal et al. (2007) observed that faecal smear examination with acid fast staining detected more goats to be positive than bacterial culture and PCR. Microscopic view of acid fast bacilli has been shown in Fig. 2. Previous studies also reported the higher prevalence of MAP in farm herds as compared to farmer’s herds (Kumar et al., 2007). Tissue samples of all the 3 goats died during study were positive by PCR (Fig. 3). In the last two decades with the advent of molecular assays like IS900 progress has been made in order to differentiate MAP strains as well as isolates for classifying them into various types and subtypes (Sohal et al., 2009). None can deny the fact that IS900 sequence is the most commonly targeted sequence for confirmation of MAP at molecular level. There are however certain concerns that the sequence may not be 100% unique to MAP.

Fig. 2: Microscopic view of acid fast bacilli (indistinguishable to MAP)

Fig. 3: MAP specific amplicons (229 bp) using IS900 PCR, Lane M: 10 bp DNA marker, Lane 1: Positive control (MAP DNA), Lane 2: Negative control (miliQ water), Lane 3-5: Tested DNA samples

Fig. 4: Gross changes showing enlargement of mesenteric and ileocecal lymph nodes

Fig. 5: Gross changes showing corrugation and thickening of intestinal mucosa

Due to homologous IS900-like elements in related mycobacterial species false positive results can occur due to cross reaction (Deb et al., 2011). Stabel (1998) reported that IS900 based PCR could be more sensitive in tissue samples in confirming diagnosis at necropsy (Singh et al., 2013a, b).

All the three goats died during the present study in Group-I were subjected to post mortem examination which revealed enlargement of mesenteric and ileo-cecal lymph nodes (Fig. 4) and thickening of ileum and ileo-cecal junction wall. In one goat slight corrugation of ileal mucosa was observed. Carcass showed depletion of fat depots. Rajukumar (1998) observed similar lesions with enlarged, edematous and congested mesenteric and ileocecal lymph nodes (Clarke, 1997; Collins et al., 2001). Corrugation of ileal mucosa (Fig. 5) observed in only one goat was as described into Ohio State University Fact Sheet for Johne’s disease, that sheep and goats may exhibit some gross and microscopic lesions not typical of those seen in cattle. Ridges and thickening of the small intestine (Fig. 5) and caecum are not always seen as is common in cattle. Gross changes present in rectum are indicated in Fig. 6 (Stabel, 1998; Sechi et al., 2005).

Fig. 6: Gross changes showing affected rectum

Storset et al. (2001) observed more number of goats with lesions in jejunum as compared to ileum while the lesions in ileum were detected in only one goat. In contrast to this lesions in jejunum were not observed in the present study. Histopathological changes in goats died of JD revealed slightly congested intestinal mucosa with infiltration of mononuclear cells consisting of lymphocytes and macrophages in lamina propria. Villi were thickened and blunt at some places. Mesenteric lymph nodes exhibited infiltration of lymphocytes and macrophages. Buergelt and Ginn (2000) as well as Munjal et al. (2005) observed similar lesions such as thickening of the intestinal villi with flat and wide tips, infiltration of lymphocytes, macrophages and epitheloid cells in lamina propria of intestine and inter-follicular area of mesenteric lymph nodes. Similar histopathological changes were also observed by Paliwal and Rajya (1982) and Rajukumar (1998).

In conclusion, the utility of multiple diagnostic tests including of conventional, serological and recent molecular tools is suggested for reaching to a confirmatory diagnosis of MAP infection in animals and studying the prevalence and epidemiology this important disease (JD). The advances in molecular diagnostic tools (Deb et al., 2011, 2013; Dhama et al., 2013a, b), inventing effective prophylactics (Dhama et al., 2008, 2013c), novel therapeutic regimens (Mahima et al., 2012; Dhama et al., 2013d, e; Tiwari et al., 2013b) along with suitable preventive and control measures (Tripathi et al., 2002; Momotani, 2012; Sweeney et al., 2012) have to be adapted and explored fully for alleviating huge economic impacts due to this significant pathogen of animals having public health concerns, particularly in the Era of One Health, One Medicine (Grant, 2005; Dhama et al., 2013e).


Authors are thankful to Council of Science and Industrial Research, New Delhi for financial assistance and Head, Animal Health Division and Director, CIRG, Makhdoom for the technical assistance.

Braun, R.K., C.D. Buergelt, R.C. Littell, S.B. Linda and J.R. Simpson, 1990. Use of an enzyme-linked immunosorbent assay to estimate prevalence of paratuberculosis in cattle of Florida. J. Am. Vet. Med. Assoc., 196: 1251-1254.
Direct Link  |  

Buergelt, C.D. and P.E. Ginn, 2000. The histopathologic diagnosis of subclinical Johne's disease in North American Bison (Bison bison). Vet. Microbiol., 77: 325-331.
CrossRef  |  Direct Link  |  

Chiodini, R.J., H.J. van Kruiningen and R.S. Merkal, 1984. Ruminant paratuberculosis (Johne's disease): The current status and future prospects. Cornell Vet., 74: 218-262.
PubMed  |  Direct Link  |  

Clarke, C.J., 1997. The pathology and pathogenesis of paratuberculosis in ruminants and other species. J. Comp. Pathol., 116: 217-261.
PubMed  |  Direct Link  |  

Cocito, C., P. Gilot, M. Coene, M. De Kesel, P. Poupart and P. Vannuffel, 1994. Paratuberculosis. J. Clin. Microbiol. Rev., 7: 328-345.
Direct Link  |  

Collins, M.T., 2002. Interpretation of a commercial bovine paratuberculosis enzyme-linked immunosorbent assay by using Likelihood Ratios. Clin. Vaccine Immunol., 9: 1367-1371.
CrossRef  |  Direct Link  |  

Collins, M.T., 2003. Paratuberculosis: Review of present knowledge. Acta. Vet. Scand., 44: 217-221.
Direct Link  |  

Collins, M.T., U. Spahr and P.M. Murphy, 2001. Ecological characteristics of Mycobacterium paratuberculosis. International Dairy Federation Report No. 362, pp: 32-40.

Deb, R. and P.P. Goswami, 2011. Coexpression of PPE 34.9 antigen of Mycobacterium avium subsp. Paratuberculosis with murine interferon gamma in HeLa cell line and study of their immunogenicity in murine model. Biotechnol. Res. Int., 10.4061/2011/632705

Deb, R., S. Chakraborty, B. Veeregowda, A.K. Verma, R. Tiwari and K. Dhama, 2013. Monoclonal antibody and its use in the diagnosis of livestock diseases. Adv. Biosci. Biotechnol., 4: 50-62.
CrossRef  |  

Deb, R., V.K. Saxena and P.P. Goswami, 2011. Diagnostic tools against Mycobacterium avium subspecies paratuberculosis infection in animals: A review. Agric. Rev., 32: 46-54.

Dhama, K., A.K. Verma, R. Tiwari, S. Chakraborty and K. Vora et al., 2013. A perspective on applications of Geographical Information System (GIS): An advanced tracking tool for disease surveillance and monitoring in veterinary epidemiology. Adv. Anim. Vet. Sci., 1: 14-24.
Direct Link  |  

Dhama, K., M. Mahendran, P.K. Gupta and A. Rai, 2008. DNA vaccines and their applications in veterinary practice: Current perspectives. Vet. Res. Commun., 32: 341-356.
CrossRef  |  Direct Link  |  

Dhama, K., M. Mahendran, R. Tiwari, S.D. Singh, D. Kumar, S. Singh and P.M. Sawant, 2011. Tuberculosis in birds: Insights into the Mycobacterium avium infections. Vet. Med. Int. 10.4061/2011/712369

Dhama, K., M.Y. Wani, R. Deb, K. Karthik and R. Tiwari et al., 2013. Plant based oral vaccines for human and animal pathogens-a new era of prophylaxis: Current and future prospective. J. Exp. Biol. Agric. Sci., Vol. 1

Dhama, K., S. Chakraborty, M.Y. Wani, R. Tiwari and R. Barathidasan, 2013. Cytokine therapy for combating animal and human diseases-A review. Res. Opin. Anim. Vet. Sci., 3: 195-208.
Direct Link  |  

Dhama, K., S. Chakraborty, Mahima, M.Y. Wani and A.K. Verma et al., 2013. Novel and emerging therapies safeguarding health of humans and their companion animals: A review. Pak. J. Biol. Sci., 16: 101-111.
CrossRef  |  Direct Link  |  

Dhama, K., S. Chakraborty, S. Kapoor, R. Tiwari and A. Kumar et al., 2013. One world, one health-veterinary perspectives. Adv. Anim. Vet. Sci., 1: 5-13.
Direct Link  |  

Ferreira, R., L.S. Fonseca and W. Lilenbaum, 2002. Agar gel immunodiffusion test (AGID) evaluation for detection of bovine paratuberculosis in Rio de Janeiro, Brazil. Lett. Applied Microbiol., 35: 173-175.
CrossRef  |  Direct Link  |  

Grant, I.R., 2005. Zoonotic potential of Mycobacterium avium ssp. paratuberculosis: The current position. J. Applied Microbiol., 98: 1282-1293.
CrossRef  |  

Green, E.P., M.L. Tizard, M.T. Moss, J. Thompson, D.J. Winterbourne, J.J. McFadden and J. Hermon-Taylor, 1989. Sequence and characteristics of IS900, an insertion element identified in a human Crohn's disease isolate of Mycobacterium paratuberculosis. Nucleic Acids Res., 17: 9063-9073.
Direct Link  |  

Kumar, P., S.V. Singh, A.K. Bhatiya, I. Sevilla and A.V. Singh et al., 2007. Juvenile Capri-Paratuberculosis (JCP) in India: Incidence and characterization by six diagnostic tests. Small Rumin. Res., 73: 45-53.
CrossRef  |  Direct Link  |  

Kurade, N.P., 1999. Studies on immunopathogenesis and enzyme linked immunosorbent assay based diagnosis in ovine paratuberculosis. Ph. D. Thesis, Deemed University, IVRI, Izatnagar.

Kurade, N.P., B.N. Tripathi, K. Rajukumar and N.S. Parihar, 2004. Sequential development of histologic lesions and their relationship with bacterial isolation, fecal shedding and immune responses during progressive stages of experimental infection of lambs with Mycobacterium avium subsp. paratuberculosis. Vet. Pathol., 41: 378-387.
CrossRef  |  Direct Link  |  

Luna, L.G., 1968. Manual of Histological Staining Methods of the Armed Forces Institute of Pathology. 3rd Edn., The Blakiston Division, McGraw-Hill Book Co., New York pp: 567-580.

Mahima, A. Rahal, R. Deb, S.K. Latheef and H.A. Samad et al., 2012. Immunomodulatory and therapeutic potentials of herbal, traditional/indigenous and ethnoveterinary medicines. Pak. J. Biol. Sci., 15: 754-774.
CrossRef  |  Direct Link  |  

Mainar-Jaime, R.C. and J.A. Vazquez-Boland, 1998. Factors associated with seroprevalence to Mycobacterium paratuberculosis in small-ruminant farms in the Madrid region (Spain). Prevent. Vet. Med., 34: 317-327.
CrossRef  |  Direct Link  |  

Manning, E.J.B. and M.T. Collins, 2001. Mycobacterium avium subsp. paratuberculosis: Pathogen, pathogenesis and diagnosis. Rev. Sci. Tech., 20: 133-150.
PubMed  |  

Molina, A., L. Morera and D. Llanes, 1991. Enzyme-linked immunosorbent assay for detection of antibodies against Mycobacterium paratuberculosis in goats. Am. J. Vet. Res., 52: 863-868.
PubMed  |  

Momotani, E., 2012. Epidemiological situation and control strategies for paratuberculosis in Japan. Jpn. J. Vet. Res., 60: S19-S29.
PubMed  |  Direct Link  |  

Munjal, S.K., B.N. Tripathi and O.P. Paliwal, 2005. Progressive immunopathological changes during early stages of experimental infection of goats with Mycobacterium avium subspecies paratuberculosis. Vet. Pathol., 42: 427-436.
PubMed  |  

Munjal, S.K., B.N. Tripathi, O.P. Paliwal, J. Boehmer and M. Homuth, 2007. Application of different methods for the diagnosis of experimental paratuberculosis in goats. Zoonosis. Public Health, 54: 140-146.
Direct Link  |  

OIE, 2000. Johne's disease in sheep and goats. Manual Ohio State University Fact Sheet for Johne's disease, pp: 30-35.

Paliwal, O.P. and B.S. Rajya, 1982. Evaluation of paratuberculosis in goats: Pathomorphological studies. Indian J. Vet. Pathol., 6: 29-34.

Pande, P.G., 1940. Paratuberculosis (Johne's disease) of cattle in Assam. Its incidence and epizootiology. Indian J. Vet. Sci., 10: 40-46.
Direct Link  |  

Paolicchi, F.A., M.J. Zumarraga, A. Gioffre, P. Zamorano and C. Morsella et al., 2003. Application of different methods for the diagnosis of paratuberculosis in a dairy cattle herd in Argentina. J. Vet. Med. B, 50: 20-26.
Direct Link  |  

Perez, V., J. Tellechea, J.M. Corpa, M. Gutierrez and J.F.G. Marin, 1999. Relation between pathologic findings and cellular immune responses in sheep with naturally acquired paratuberculosis. Am. J. Vet. Res., 60: 123-127.
PubMed  |  Direct Link  |  

Perez, V., J.F. Garcia Marin and J.J. Badiola, 1996. Description and classification of different types of lesion associated with natural paratuberculosis infection in sheep. J. Comp. Pathol., 114: 107-122.
PubMed  |  

Rajukumar, K., 1998. Pathology and diagnosis of caprineparatuberculosis. M.V.Sc. Thesis, Deemed Unjversity, IVRI, Izatnagar.

Sechi, L.A., A.M. Scanu, P. Molicotti, S. Cannas and M. Mura et al., 2005. Detection and Isolation of Mycobacterium avium subspecies paratuberculosis from intestinal mucosal biopsies of patients with and without crohn's disease in sardiniaMAP in crohn's disease. Am. J. Gastroenterol., 100: 1529-1536.
CrossRef  |  Direct Link  |  

Sechi, L.A., M. Manuela, F. Tanda, Li. Amelia, S. Antonello, F. Giovanni and Z. Stefania, 2001. Identification of Mycobacterium avium subsp. paratuberculosis in biopsy specimens from patients with Crohn;s disease identified by in situ hybridization. J. Clin. Microbiol., 39: 4514-4517.
CrossRef  |  Direct Link  |  

Shankar, H., S.V. Singh, P.K. Singh, A.V. Singh, J.S. Sohal and R.J. Greenstein, 2010. Presence, characterization and genotype profiles of Mycobacterium avium subspecies paratuberculosis from unpasteurized individual and pooled milk, commercial pasteurized milk and milk products in India by culture, PCR and PCR-REA methods. Int. J. Infect. Dis., 14: e121-e126.
CrossRef  |  PubMed  |  Direct Link  |  

Sherman, D.M., R.J. Markham and F. Bates, 1984. Agar gel immunodiffusion test for diagnosis of clinical paratuberculosis in cattle. J. Am. Vet. Med. Assoc., 185: 179-182.
PubMed  |  

Singh, A.V., S.V. Singh, J.S. Sohal and P.K. Singh, 2009. Comparative potential of modified indigenous, indigenous and commercial ELISA kits for diagnosis of Mycobacterium avium subspecies paratuberculosis in goat and sheep. Indian J. Exp. Biol., 47: 379-382.
PubMed  |  Direct Link  |  

Singh, A.V., S.V. Singh, P.K. Singh, J.S. Sohal and N. Swain et al., 2009. Multiple tests based prevalence estimates of Mycobacterium avium subspecies Paratuberculosis infection in elite farms of goats and sheep. J. Small Rumin., 15: 178-182.
Direct Link  |  

Singh, N., S.V. Singh, V.K. Gupta, V.D. Sharma, R.K. Sharma and V.M. Katoch, 1996. Isolation and identification of Mycobacterium paratuberculosis from naturally infected goatherds in India. Indian J. Vet. Path., 20: 104-108.

Singh, P.K., S.V. Singh, H. Kumar, J.S. Sohal and A.V. Singh, 2010. Diagnostic application of IS900 PCR using blood as a source sample for the detection of Mycobacterium avium subspecies paratuberculosis in early and subclinical cases of caprine paratuberculosis. Vet. Med. Int., 10.4061/2010/748621

Singh, S.V., A.V. Singh, A. Kumar, P.K. Singh and R. Deb et al., 2013. Survival mechanisms of Mycobacterium avium subspecies paratuberculosis within host species and in the environment: A review. Nat. Sci., 5: 710-723.
CrossRef  |  Direct Link  |  

Singh, S.V., A.V. Singh, P.K. Singh, J.S. Sohal and N.P. Singh, 2007. Evaluation of an indigenous ELISA for diagnosis of Johne's disease and its comparison with commercial kits. Indian J. Microbiol., 47: 251-258.
CrossRef  |  PubMed  |  Direct Link  |  

Singh, S.V., A.V. Singh, P.K. Singh, J.S. Sohal and Y. Vikram et al., 2007. Sero-prevalence of Mycobacterium avium subspecies paratuberculosis infection in Barbari and Sirohi breeds of goats in their respective home tracts in semi-arid regions of North India. Haryana Vet., 46: 104-106.

Singh, S.V., A.V. Singh, S. Gupta, A.S. Rajindran and N. Swain et al., 2012. Interspecies sharing of indian bison type a novel predominant genotype of Mycobacterium avium subspecies Paratuberculosis between naturally infected and endemic flocks of bharat merino sheep and a colony of rabbits (Oryctolagus cuniculus) raised on the same ecosystem in South India. Res. Rev. J. Life Sci., 2: 1-8.
Direct Link  |  

Singh, S.V., K. Dhama, K.K. Chaubey, N. Kumar and P.K. Singh et al., 2013. Impact of host genetics on susceptibility and resistance to Mycobacterium avium subspecies Paratuberculosis infection in domestic ruminants. Pak. J. Biol. Sci., 16: 251-266.
CrossRef  |  Direct Link  |  

Sockett, D.C., D.J. Carr and M.T. Collins, 1992. Evaluation of conventional and radiometric fecal culture and a commercial DNA probe for diagnosis of Mycobacterium paratuberculosis infections in cattle. Canadian J. Vet. Res., 56: 148-153.
Direct Link  |  

Sohal, J.S., S.V. Singh, A.V. Singh and P.K. Singh, 2009. Strain diversity within Mycobacterium avium subspecies paratuberculosis: A Review. Indian J. Exper. Biol., 48: 7-16.
PubMed  |  Direct Link  |  

Stabel, J.R., 1998. Johne's disease: A hidden threat. J. Dairy Sci., 81: 283-288.
CrossRef  |  PubMed  |  Direct Link  |  

Stewart, D.J., J.A. Vaughan, P.L. Stiles, P.J. Noske and M.L.V. Tizard et al., 2006. A long-term study in Angora goats experimentally infected with Mycobacterium avium subsp. paratuberculosis: Clinical disease, faecal culture and immunological studies. Vet. Microbiol., 113: 13-24.
CrossRef  |  Direct Link  |  

Storset, A.K., H.J. Hasvold, M. Valheim, H. Brun-Hansen and G. Berntsen et al., 2001. Subclinical paratuberculosis in goats following experimental infection: An immunological and microbiological study. Vet. Immunol. Unopathol., 80: 271-287.
CrossRef  |  PubMed  |  Direct Link  |  

Sweeney, R.W., D.E. Jones, P. Habecker and P. Scott, 1998. Interferon-gamma and interleukin 4 gene expression in cows infected with Mycobacterium paratuberculosis. Am. J. Vet. Res., 59: 842-847.
PubMed  |  Direct Link  |  

Sweeney, R.W., M.T. Collins, A.P. Koets, S.M. McGuirk and A.J. Roussel, 2012. Paratuberculosis (Johne's Disease) in cattle and other susceptible species. J. Vet. Internal Med., 26: 1239-1250.
CrossRef  |  PubMed  |  Direct Link  |  

Sweeney, R.W., R.H. Whitlock, C.L. Buckley and P.A. Spencer, 1995. Evaluation of a commercial enzyme-linked immunosorbent assay for the diagnosis of paratuberculosis in dairy cattle. J. Vet. Diagn. Invest., 7: 488-493.
CrossRef  |  Direct Link  |  

Tiwari, R., K. Dhama, S. Chakraborty, A. Kumar, A. Rahal and S. Kapoor, 2013. Bacteriophage therapy for safeguarding animal and human health: A review. Pak. J. Biol. Sci., (In Press).

Tiwari, R., S. Chakraborty, K. Dhama, S. Rajagunalan and S.V. Singh, 2013. Antibiotic resistance-an emerging health problem: Causes, worries, challenges and solutions: A review. Int. J. Curr. Res., 5: 1880-1892.
Direct Link  |  

Tripathi, B.N., S. Periasamy, O.P. Paliwal and N. Singh, 2006. Comparison of IS900 tissue PCR, bacterial culture, Johnin and serological tests for diagnosis of naturally occurring paratuberculosis in goats. Vet. Microbiol., 116: 129-137.
CrossRef  |  Direct Link  |  

Tripathi, B.N., S.K. Munjal and O.P. Paliwal, 2002. An overview of paratuberculosis (Johne's disease) in animals. Indian J. Vet. Pathol., 26: 1-10.

Van Embden, J.D.A., M.D. Cave, J.T. Crawford, J.W. Dale and K.D. Eisenach et al., 1993. Strain identification of Mycobacterium tuberculosis by DNA fingerprinting: Recommendations for a standardized methodology. J. Clin. Microbiol., 31: 406-409.
PubMed  |  

Vary, P.H., P.R. Andersen, E. Green, J. Hermon-Taylor and J.J. McFadden, 1990. Use of highly specific DNA probes and the polymerase chain reaction to detect Mycobacterium paratuberculosis in Johnes disease. J. Clin. Microbiol., 28: 933-937.
PubMed  |  

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