Diagnosis of Bovine Tuberculosis

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This entry is adapted from the peer-reviewed paper 10.3390/pathogens11070715

Bovine tuberculosis is a serious infectious disease affecting a wide range of domesticated and wild animals, representing a worldwide economic and public health burden. The disease is caused by Mycobacterium bovis and infrequently by other pathogenic mycobacteria. The problem of bovine tuberculosis is complicated when the infection is associated with multidrug and extensively drug resistant M. bovis. Many techniques are used for early diagnosis of bovine tuberculosis, either being antemortem or postmortem, each with its diagnostic merits as well as limitations. Antemortem techniques depend either on cellular or on humoral immune responses, while postmortem diagnosis depends on adequate visual inspection, palpation, and subsequent diagnostic procedures such as bacterial isolation, characteristic histopathology, and PCR to reach the final diagnosis.

bovine tuberculosis Diagnosis Laboratory

1. Field Diagnosis of Bovine Tuberculosis (bTB)

1.1. Tuberculin Test

Tuberculin skin test is the recommended standard procedure for ante-mortem diagnosis of Bovine Tuberculosis (bTB), which evaluates a delayed-type hypersensitivity reaction in sensitized cattle following an intradermal injection of purified protein derivate (PPD) tuberculin from mycobacteria, by measuring changes in skin thickness after 72 h from injection. In Egypt, as well as many other countries, PPD is injected in the mid-cervical region (cervical intradermal test), whilst in other countries as North America it is applied in the caudal fold of the tail (caudal fold test) ^[1]. The PPD injection stimulates the CMI and sensitizes T cells by prior infection which accumulate at the injection site and release lymphokines causing local vasodilatation, edema, fibrin deposition, aggregation of other inflammatory cells, and ultimately forming local skin swelling ^[2].

There are two commonly used types of the intradermal tuberculin test. The single intradermal (SID), in which PPD of M. bovis is used, despite its wide usage, high availability, and low costs, fails to differentiate between bTB infected cattle and others sensitized with M. avium complex or environmental mycobacteria. The second type is the comparative intradermal tuberculin test (CITT), in which both bovine and avian PPDs are injected simultaneously to increase the specificity of TST ^[3]. In Egypt, CITT is not used by general organization of veterinary services (GOVs) due to the lack of financial support. However, it is used only privately by owners of some herds ^[4].

OIE (2009) ^[5] define the standard TST procedures. (1) Clipping and shaving of the injected site. (2) Measuring a fold of skin of clipped area with a caliper. (3) A short needle with bevel edge in graduated tuberculin syringe is inserted obliquely into skin and the dose is injected. The recommended dose of bovine PPD must not be lower than 2000 IU. In cattle with diminished allergic sensitivity, a higher dose of PPD is needed, and in national eradication campaigns, doses of up to 5000 IU are recommended. (4) Palpating of a pea-like swelling at the injected area is a sign of correct intradermal injection. (5) The distance between two injections must be 12–15 cm distance. (6) The skin fold thickness is remeasured after 72 h where the same operator should measure the two readings. The positive bovine response is considered when the swelling is >4 mm, while <3 mm is considered negative, and between 3 and 4 mm is suspicious ^[6].

Byrne et al. found a strong positive relationship between the reaction size of injected PPDs and the severity of PM lesions and suggested that increased reaction size is indicative of progressive lesions.

Despite its wide use worldwide, TST has several drawbacks: (1) Complexity. (2) The interpretation is subjective and differs between operators. (3) False positive reactions when the animals were TST positive but there is no PM lesions nor positive culture, known as Non-Visible Lesions (NVL). They are given by cross reactivity to other non-virulent mycobacteria, avian TB, human TB, Johne’s disease (JD), or sensitized by other allergens as Nocardia farcinicus causing non-specific responses. Additionally, they occur in early stages of the disease, where TB granulomas are too small or infrequent to be seen during PM examination, or in infected animals during the latency in which infection with M. bovis, but without disease. (4) False negative reactions and those given during the late stage of infection, particularly in severe and generalized cases where the animals are non-responsive for TST and known as the State of Anergy. Also, the false negative reactions are given in early cases until 3–6 weeks post-infection which is known as the pre-allergic period. Further false negatives include animals desensitized by PPD administration during the preceding 8 to 60 days, old cattle, early post-partum cows (postparturient desensitization because of the general immunologic hyporeactivity), and low potency tuberculin, subcutaneous injection (rather than intradermal), or bacterial contamination of the tuberculin ^[7]^[8]^[9].

There are various factors affecting the sensitivity and specificity of TST, that range from 55–99%, such as potency, purity, dosage, and biological activity of the PPD, as well as the inoculation site, misreading of the results and the genetic background of the animal ^[3]^[9]. In addition, some co-infections affect the performance of TST particularly, the infection with M. avium subspecies paratuberculosis (MAP), the causative agent of Johne’s Disease, such cross-reactivity lead to misdiagnosis as MAP shares structural proteins and virulence factors with M. bovis ^[10]. In addition, co-infection with bovine viral diarrhea (BVD) virus has conflicting effects in previous studies; it is suggested to suppress the immunological response to PPD or cause rapid progression of bTB or even to have no significant effect ^[11]. Furthermore, co-infection with liver fluke, Fasciola hepatica and Fasciola gigantica, may affect the accuracy of bTB but the direction of the effect differs among studies ^[12]. It is suggested that liver fluke infection may suppress the M. bovis infection ^[11]. Howell et al. examined the evidence to determine the effect of liver fluke infection on four outcomes relevant to bTB diagnosis: TST, IGRA, lesion detection, and bacterial culture ^[12]. The study supported the hypothesis that liver-fluke-infected animals are likely to have a reduced response to both TST and IGRA tests and fewer bacteria recovered/cultured from their lesions. Furthermore, technical and procedural mistakes during TST application, related to measuring of skin thickness, storage and injection of PPD, and interpretation of test results, negatively affected the true detection of bTB ^[13].

Bovine PPD is a mixture of protein, carbohydrates, and lipids obtained from M. bovis AN5 and some of these components are present in nonpathogenic environmental mycobacteria ^[14]. Therefore, and for improvement sensitivity and specificity of TST, some trials for using some antigens instead of PPDs were performed; Parlane et al. displayed four mycobacterial proteins (ESAT-6, CFP10, Rv3615c, and Rv3020c) at high density on bacteria-produced polyester inclusions (biobeads) and concluded that their use should allow the development of a highly sensitive, specific, and cost-effective skin test for diagnosis of bTB ^[15].

Two approaches have been used globally at the herd level to overcome the limited sensitivity of TST in case of high prevalence settings. The first is the use of in vitro ancillary tests to maximize the detected number of infected animals and the other is the depopulation of the herds whose reactors are detected. Nevertheless, high costs and economic and social implications make whole herd depopulation difficult and inapplicable, particularly in endemic setting ^[16].

European Bison (Bison bonasus) is extremely sensitive to mycobacterial infection and its ongoing restitution requires the improvement of ante-mortem diagnostic methods such as TST and IGRA ^[17].

1.2. Ante-Mortem Examination (Clinical Signs) of bTB

The clinical signs of bTB vary depending on several factors: the sites of localization of infection, infectious dose, virulence, state of immune competence of the host, and external influences ^[18]. The incubation period ranges between 2 months, at minimum, and several years ^[19]. Most cattle that are infected do not develop clinical signs, but when present they are extremely variable and often nonspecific ^[20]. Because the disease is always progressive, there is the constant underlying toxemia, which causes weakness, debility, and the eventual death of the animal ^[8].

Pulmonary involvement is characterized by a chronic moist cough, thoracic abnormalities on auscultation heard especially in the advanced stages when much lung tissue has been destroyed, dyspnea with increased rate and depth of respiration become apparent. Lymph node enlargement coupled with chronic respiratory disease may result in a higher index of suspicion. Enlargement of bronchial LNs may cause dyspnea because of constriction of air passages, while retropharyngeal LN involvement may cause either respiratory signs and noisy breathing, or dysphagia and eructation. Meanwhile, the enlargement of the mediastinal LN is commonly associated with recurrent and then persistent ruminal tympany. Rarely, tuberculous ulcers of the small intestine can cause diarrhea. The reproductive disorders associated with bTB are uncommon; tuberculous metritis causing infertility sometimes with chronic purulent discharges, vaginitis, and rare cases of tuberculous orchitis. Tuberculous mastitis is of a great significance because of the public health danger, it is characterized by marked induration and hypertrophy, which usually develops first in the upper part of the udder, with supra-mammary LN enlargement. In case of miliary TB, some cows with extensive lesions are clinically normal, but in most cases progressive emaciation unassociated with other signs occurs. A capricious appetite and fluctuating temperature are also commonly associated with the disease ^[8]^[20].

1.3. Postmortem Examination of bTB

Postmortem examination is a cornerstone for bTB control programs in endemic areas to detect the infection either in routinely slaughtered animals or in tuberculin test reactors ^[21]. Despite the wide availability of tests for the identification of M. bovis infection at the herd level, the diagnosis of bTB is often difficult due to the scarcity of diagnostic tests that fulfill all the essential criteria necessary for the identification of infected animals. It is noteworthy that almost the 20–30% of the new bTB cases are first diagnosed during postmortem inspection at the slaughterhouse in cattle intended for human consumption ^[21]^[22].

Bovine TB is characterized by formation of granulomatous nodules called tubercles which are circumscribed yellowish inflammatory nodules approximately 2–20 mm in diameter that are more or less encapsulated by connective tissue and often contain central caseous necrosis and mineralization. Tubercles are found in the LNs, particularly bronchial, retropharyngeal, and mediastinal nodes. They are also common in the lung, spleen, liver, heart, kidney, and surfaces of the body cavities ^[23]^[24]. Lesions may occasionally be found on pleural sac and in mammary tissues, some of them with cheese-like foci on cut surfaces. In general, some lesions appear as abscesses containing yellowish pus ^[25]. The pus has a variable color range from a characteristic creamy to orange, and consistencies from thick cream to crumbly cheese. In addition, lesion size may be microscopic or large enough to involve the greater part of the whole organ or tissue ^[8]^[26].

In lungs of extensively affected cases, diffuse pus is evident and may spread to cause suppurative bronchopneumonia as well as adhesion to the pleural cavity due to fibrinous inflammation. In addition, the presence of bronchopneumonia or hyperemia around pulmonary lesions is highly suggestive of active disease ^[8]^[27]. The chronic lesions become considerably enlarged, nodular, and contain thick, yellow to orange, caseous material, often calcified and surrounded by a thick fibrous capsule ^[8]. In disseminated cases, the miliary tuberculosis form is characterized by a large number of small grey to white yellowish caseous foci resembling millet seeds without clear-cut delimitations that are found throughout the lung and other organs. In addition, generalization to the serosal surfaces, especially the pleura, pericardium, or peritoneum, may occur and is characterized by multiple small tubercles of approximately 0.5 to 1 cm in diameter that resemble pearls. Lesions are sometimes found in the female genitalia but are rare in the male genitalia ^[23]^[24].

Interestingly, not all animals with bTB-like lesions would be detected at PM examination even with perfect examination techniques, because some lesions are invisible to the naked eye. The primary complex is most frequently located in the lower parts of the respiratory tract, 70% of these lesions may only be found during careful examination and dissection of the lungs into thin sections. Not all infected animals have lesions at the time of slaughter, as the existence of bTB lesions is directly related to the interplay between the host’s defense mechanism and mycobacterial virulence factors ^[21]^[23]. Moreover, it is important to point out other diseases whose lesions may resemble tubercles are common in other animal species ^[28]^[29]^[30]. Furthermore, most Mycobacterium-associated lesions are indistinguishable from mixed infections or lesions caused by other bacteria such as Rhodococcus equi. In cattle, pigs, and wild boars, R. equi infection is mainly associated with tuberculous-like lesions in lymph nodes. R. equi was isolated from cattle and American bison with purulent lesions suspected of Mycobacterium spp. infection. In pigs, R. equi, not the previously suspected Mycobacterium spp., is the primary causal agent of lymphadenitis.

2. Laboratory Diagnosis

2.1. IFN-γ Release Assay (IGRA)

IFN-γ release assay is deemed the main ancillary test in bTB diagnosis and is usually used in parallel with TST ^[31]. It relies on in-vitro measurement of IFN-γ cytokine and has many advantages over other antemortem diagnostic tests, it being more sensitive, faster, and requiring one farm visit ^[32]. Additionally, it has one major advantage over TST, being able to detect more earlier infections than TST where a hypersensitivity reaction against PPD has developed between 3–6 weeks post-infection ^[14], while IGRA can detect the infection as early as 14 days after infection ^[33]. Thus, it can detect a substantial proportion of infected animals that escape detection by TST ^[9]. Moreover, it is also used to diagnose tuberculosis-similar diseases ^[34], and there are no limitations for retesting by IGRA because it, in contrast to TST, is an in vitro test.

Many previous studies reported that the prior administration of TST causes either a booster or a drop in IFN-γ release. These discrepancies are likely to be related to the variable conditions and type of skin test in each study ^[9]^[35]. Because TST may cause an apparent increase of IFN-γ production, it has been suggested to perform IGRA after 33 days to minimize its effect on the results ^[35]. In addition, Elsohaby et al. performed IGRA 45 days after TST ^[36]. In contrast, Clarke et al. studied the effect of timing of blood collection for IGRAs relative to the TST application in African buffaloes and recommended that collection of blood samples prior to or at the time of TST had a significance in the detection of a greater number of positive buffaloes than their collection three days after TST ^[37]. In the same line, parallel using of TST and IGRA maximized the detection of infected animal and was able to detect all infected buffaloes according to ^[38]. Additionally, De la Rua-Domenech et al. mentioned that blood samples can be taken as early as 3 days post-TST without affecting the results of the assay ^[9].

Abdellrazeq et al. pointed out, for the first time, cut-off criteria to optimize IGRA as a routine ancillary test for diagnosis of bTB in Egypt ^[4]. Also, Elsohaby et al. in Egypt, estimated the sensitivity and specificity of IGRA, PCR, and mycobacterial culture for detection of M. bovis in blood and milk, and recorded a higher sensitivity of IGRA than PCR and culture, and recommended the use of IGRA in the Egyptian bTB eradication program ^[36].

Despite its advantages, IGRA has high logistical demands (must be cultured within 24 h after blood sampling), and of high costs ^[39]. IGRA also has some limitations arising from using of PPD as antigen due to cross reactivity to environmental mycobacteria, leading several studies to develop multiple-antigen cocktails (Mb1762c, Mb2054c, Mb2057c, and Mb2660c) to increase the sensitivity of IGRA when supplemented to PPDs ^[40]. In addition, use of ESAT6 and CFP10 antigens instead of PPD in IGRA increased the ability to identify bTB infected animals and to distinguish them from NTM-exposed or BCG-vaccinated animals ^[6].

Novel biomarkers to distinguish between the healthy and infected animals are still urgently needed, particularly when TST and IGRA fail to detect the infection ^[3]. Hence, several biomarkers have been used in blood-based TB tests as interleukin (IL)-1β, IL-2, IL-17, IL-21, IL-13, IL-22, chemokine C-X-C motif ligand 9 (CXCL9), and CXCL10. Many of these cytokines are suggested for use as diagnostic biomarkers of M. bovis infection in cattle ^[41].

Both TST and IGRA detect the early bTB infection as they depend on the measurements of pathogen specific CMI responses ^[42]. At advanced stages of the disease, CMI responses decrease in parallel with increasing of the humoral immune response. Thus, TST and IGRA can give false negative results at the late stages ^[9]. Therefore, diagnosis of anergic animals, that show no response to CMI-based tests, is critical, because these animals could relapse and cause future outbreaks ^[43]. Thus, serological tests such as ELISA at the late stages of the disease are highlighted, especially with the adding of secreted proteins as MPB70 and MPB83 that are notably released by M. bovis in large amounts during the late stages of the disease ^[14]^[39].

2.2. ELISA

Various factors affect the sensitivity and specificity of serological tests for diagnosis of bTB such as the stage of the disease, immune status of the animal, type of antigen used, and previous exposure to bovine tuberculin ^[44]. The sensitivity of antibodies-based ELISA is higher when evaluated in animals at later stages of the disease with gross lesions and in most infective animals ^[45]. In addition, they offer some advantages over CMI-testing, such as relative ease of sample collection, greater practicality, cost-effectiveness, and their ability to detect anergic animals ^[46]. On the other hand, others consider ELISA tests to be of a lower accuracy employ them less frequently than CMI-based tests for diagnosis of bTB due to their lower sensitivity and highly variable overall test performance ^[46]^[47]. Despite their precise role not being well understood, many factors participate in the variable performance of ELISA tests, such as geographical location, stage of infection and exposure to and diversity of NTMs ^[46].

TST is known to boost the antibody response; thus, the use of ELISA without skin tests would further reduce their sensitivity ^[47]. Testing of different sampling times is required to evaluate the best time to collect serum samples after PPD injection based on the increased sensitivity of the serological test ^[14]. Thus, several recent studies used ELISA tests to complement a prior TST; Casal et al. evaluated two ELISAs on sera collected prior to, and 3 days and 15 days after PPD injection, and reported the highest level of detected animals in samples collected after 15 days after TST, taking advantage of the anamnestic effect (increased serological response after performance of TST leading to an improvement of the sensitivity of the used technique) ^[45]. In addition, Fontana et al. validated a multi-antigen ELISA comprising five antigens (ESAT6, CFP10, PPD-B, MPB70, and MPB83) in sera collected 15–20 days after a single TST and demonstrated 74.2% and 94.9% of sensitivity and specificity, respectively ^[43]. Also, Souza et al. applied a recombinant chimera ELISA antigen of ESAT-6, MPB70, and MPB83 on sera obtained 7 days after a comparative TST, the sensitivity and specificity of the ELISA were 79.5%, and 75.5%, respectively ^[48]. More recently, Griffa et al. used an antigenic mixture from a total extract of the reference strain AN5 and were able to confirm the M. bovis infection of 83.7% of animals that were ELISA positive 15–17 days after a negative TST, by histopathology and PCR ^[49]. In the same study, the specificity was 95.95% and the authors suggested the detection of antibodies of M. bovis within weeks after TST as a rapid and inexpensive way to improve bTB control. On the other hand, Casal et al. took the serum samples before the injection of PPD and evaluated the sensitivity of three different ELISAs in addition to TST and two IGRA tests for the diagnosis of bTB in cattle and concluded that in vitro diagnostic techniques maximized the detection of bTB infected animals ^[50]. The authors suggested the parallel use of cellular and humoral-based tests in high prevalence setting conditions to accelerate bTB eradication because the antibodies-based tests significantly improved the sensitivity of cellular based tests up to 98.2%. Contrarily, McCallan et al. took the samples prior to TST to assess the utility of three serological tests, and their study reported that serological tests were of limited advantage when used in parallel with TST and IGRA, because the serological tests disclosed only about 3% of positive animals whilst TST and IGRA disclosed 13% and 40% of positive animals, respectively ^[51].

Several studies highlighted the great significance of MPB83 protein as a diagnostic antigen in serological tests and recommended its use because of the highest and earlier response that was triggered against it ^[14]. Waters et al. reported that antibodies responses against MPB70 and MPB83 reach their peak 2 weeks after injection of PPD then begin to wane 1–2 months post TST, and can be further increased after re-injection of PPD ^[52].

IDDEX™ ELISA is a commercial kit recognized by OIE for detection of bTB infection in blood and milk serum samples depending on detection of antibodies against MPB70 and MPB83 antigens. It is used in several studies: Al-Fattli in Iraq used it as a screening test and reported a seroprevalence of 20.16% and 15.12% in blood and milk of lactating cows, respectively ^[53]. However, Soares Filho et al. recommended that IDDEX™ ELISA cannot be used as a single test for PM diagnosis of bTB because of its low sensitivity despite the test being able to detect eight positive samples that were negative on RT-PCR and culture ^[44]. The authors collected blood and obtained serum aliquots from the brachial vein venous blood of the half carcass from which the respective tissue and organ samples were obtained. Trost et al. reported a wide variation in sensitivity of IDDEX™ ELISA by geographical distribution, where it was 9%, 45%, and 77% from bovine serum samples in Mexico, the United States, and the United Kingdom, respectively ^[54].

Other serological tests are used in several studies, such as rapid lateral-flow test for detection of antibodies against M. bovis mpb70 antigen. Elsohaby et al. used it in conjugation with TB-Feron test (a type of IGRA test) as ancillary tests and both were able to reduce a significant number of false positive TST slaughtered cows ^[55].

All the previous tests, TST, IGRA, and ELISA, have wide various limitations related to their sensitivity and specificity, thus, parallel use of more than one test and focusing on finding new cocktail mycobacterial antigens offer substantial advantages for maximizing the detection of infected animals ^[6]^[14]^[40].

2.3. Mycobacterial Culture

Isolation and identification of the mycobacteria is still the international gold standard test in diagnosis of bTB ^[5]. It can take up to three months due to the slow growth rate of MBTC ^[56].

The most frequently media used for mycobacterial growth are Lowenstein–Jensen buffered egg potato medium, Middle brook 7H10, Middle brook 7H11, and Dubos Oleic-Albumin agar. Unlike Mtb and M. avium that grow well on glycerol containing media and known as eugenic, M. bovis has a sparse thin growth on them and is called dysgenic, but it grows well on pyruvate-containing media without glycerol ^[57]. This dysgenic growth of M. bovis in the presence of glycerol is due to lack of pyruvate kinase enzyme ^[58]. The growth of moist, white, flat, and friable colonies is indicative for the primary cultures of M. bovis on Lowenstein–Jensen slants ^[59]. Zihel–Neelsen staining should be performed to confirm the presence of acid-fast bacilli. Despite being rapid and sensitive, Zihel–Neelsen lacks specificity and cannot differentiate between members of MBTC ^[60].

A range of pre-culture treatments, such as decontamination, homogenization, and concentration are conducted before inoculation on the media to facilitate the recovery of M. bovis ^[61]. The most traditional decontamination method used to isolate M. bovis from bovine tissues is the Petroff method in which 4% NaOH solution is used as a decontaminant (OIE 2014).

Mohamed et al., compared between BACTEC MGIT 960TM as a fully automated liquid-medium system and the conventional culture using Lowenstein–Jensen media for isolation of mycobacteria and found that automated system was more sensitive, faster, and revealed a higher recovery rate of mycobacteria than Lowenstein–Jensen media ^[60].

Issa et al. in Brazil, compared three decontaminants used in mycobacterial decontamination: 2% sodium hydroxide (NaOH), 0.75% hexadecyl pyridinium chloride (HPC), and 5% sulphuric acid (H₂SO₄) ^[62]. In addition, they evaluated four mycobacterial media: Middlebrook 7H11 with additives and OADC (oleic acid, albumin, dextrose, and catalase) supplement A (7H11-A), Middlebrook 7H11 with another supplement trademark (7H11-B), tuberculosis blood agar (B83), and Stonebrink’s medium.

For achieving the best results, Soares Filho et al. recommended two sampling protocols for both PCR and isolation ^[44]. For PCR, the tissue was collected from the center of the caseous lesion, where more genetic material and fewer viable bacteria are expected to be found whilst, for isolation, collection was done at the border between the healthy tissue and the lesion, where more viable bacteria, but less bacterial genetic material, would be expected.

Despite mycobacterial culture usually being regarded as the golden standard, it is time consuming and prolonged for several months, risky, laborious, with a high level of tissue sample contamination and decreased number of viable mycobacteria due to the decontamination methods ^[44]^[61]^[62]. In addition, Albernaz et al. reported a low sensitivity of bacterial culture and stated that it is not recommended as a routine complementary test for diagnosis of bTB in buffaloes ^[63]. The major limitation of mycobacterial culture is being confined to PM lesion samples, some studies suggest the use of nasal swabs as an alternative method ^[39]. However, Mayer et al. stated that RT-PCR from nasal swabs is not suitable for in vivo diagnosis of bTB ^[64].

2.4. Histopathology as a Diagnostic Method of bTB

Canal et al. described the characteristic histopathological pictures of tuberculous lesions that were classified microscopically into four stages (stage I, II, III, and IV) ^[65]. Stage I, in which irregular epithelioid macrophages, dispersed lymphocytes, and few Langhans-type multinucleated giant cells are displayed. Stage II granulomas exhibited limited necrosis with neutrophils, lymphocytes, macrophages in addition to few fibroblasts and Langhans-type cells. Stage III granulomas exhibited epithelioid and Langhans-type giant cells in the peripheral areas of the central caseous necrosis, with central calcification. Near the fibrous capsule, the inflammatory cell population consisted of macrophages, lymphocytes, and scattered neutrophils. Stage IV granulomas exhibited mostly necrosis and mineralization. A large fibrous capsule was evident, and this capsule shaped an irregular area of large necrosis and mineralization. Evidence of thick encapsulated lesions is suggestive for lower dissemination and an active anti M. bovis immune response.

Despite high sensitivity, histopathology lacks specificity; McKinley et al. described a histopathological profile with encapsulation of granuloma, presence of Langhan’s cells, sometimes in association with epithelioid cells, lymphocytes, or neutrophils in tuberculous-like lesions ^[66]. However, neither M. bovis nor member of MBTC were detected either by molecular methods or cultivation over 3 months, but NTM and Actinomycetales were identified in the lesions.

2.5. Molecular Diagnosis

PCR techniques are widely used for the diagnosis of bTB and have several advantages; they are quick, applied within a few hours which means rapid diagnosis and efficient control, overcome the lack of specificity of other traditional tests such as histopathology, and are able to identify the mycobacteria either from culture or clinical specimens ^[32]^[67]. Moreover, several PCR techniques targeting numerous genes and regions can be used for differentiation between members of MBTC, such as ESAT-6 and CFP-10, which are protein products of esxA and esxB genes, respectively ^[68], atpE and lpqT ^[69], regions of difference (RD 1, 4, 9, 12) ^[69]^[70], RvD1-Rv2031c ^[40], and insertion sequences (IS) as IS1081 ^[44] and IS6110. Numerous studies targeted the insertion sequence region IS6110 as it is found in all members of MBTC ^[67]^[68]^[71].

Thacker et al. reported a high specificity of TaqMan Real-Time PCR targeting IS6110 gene as it was able to detect 5 pg/μL of M. bovis specific DNA or even smaller quantities in tissue samples ^[71]. Despite Soares Filho et al. reporting a low sensitivity of the PCR technique as a PM diagnostic method of suggestive tuberculous lesions, the authors recommended the usage of PCR in situations of high prevalence or in parallel with other tests such as ELISA because it is a quick, safer, and relatively less expensive technique ^[44]. On the contrary, Algammal et al. reported a higher sensitivity of PCR, over 85%, compared to other PM diagnostic methods ^[32].

Several other reports detected lower sensitivity and specificity of PCR. The sensitivity of the molecular studies varies from 50% to more than 80% depending on the study and the employed methods ^[72]. The complex mycobacterial cell wall, presence of the bacterial cell within granulomas and the presence of PCR inhibitors and subsequent failure of DNA extraction are obstacles facing the PCR diagnosis of MBTC organisms ^[73]^[74].

The varying results of PCR technique performance are mainly because of technical differences in the setting up of assays, particularly during DNA extraction from lesions ^[44]^[74], and their sensitivity is conditional on sensitivity of necropsy and volume of DNA ^[75]. Further, contamination of the PCR reaction and the presence of environmental bacteria can prompt false positives and cause insufficient specificity ^[71]. Differences of the PCR primers used ^[47] and the presence of inhibitory substances in samples or reagents can also cause reduced sensitivity ^[74].

There is a significant variation of sensitivity of PCR assays related to the DNA extraction method used and there is not a definitive view for the best method of DNA extraction from bovine tissues ^[76]. Hence, several studies compared DNA extraction protocols; Ikuta et al. compared three protocols and reported 46.6%, 50%, and 100% positive samples of the three protocols from the same samples ^[74]. In addition, Moura et al. evaluated nine DNA extraction methods, using nine commercial kits, reported various results between them and concluded that DNA extraction kit deeply influences the diagnostic sensitivity of bTB in bovine tissue samples ^[77].

Despite their advantages, PCR techniques have several limitations; limited to the PM diagnosis ^[44], not specific for pathogen identification, being restricted only to members of MBTC or M. avium complexes ^[78]. In addition, reduced sensitivity of some PCR assays due to inhibitory substances in samples or reagents, during DNA extraction, as well as lower amounts of DNA ^[26]^[74]. Helmy et al. reported that RT-PCR was more sensitive and specific than conventional and multiplex PCR, less manipulated and possessed low risk of cross-contamination ^[79]. In addition, Algammal et al. reported that RT-PCR is the most sensitive rapid diagnostic test for detection of M. bovis from tissues and provides higher positive values than culturing ^[32]. However, other studies confirmed the usefulness of combination between RT-PCR and conventional PCR for rapid identification and discrimination between members of MBTC ^[69].

Besides tissue samples, many PCR assays had been performed on blood and milk samples such as the Mataragka et al. study, which illustrated that PCR can be used as an early and sensitive indicator method to detect infection in pooled milk samples collected from the aged animals of a dairy farm, which could support TST monitoring and improve bTB control ^[80]. In addition, Brahma et al. reported a similar sensitivity of PCR targeting CFP-10 protein to that of IGRA and concluded that it may be used as a fast, alternative method for bTB diagnosis from blood samples ^[81]. Elsohaby et al. reported a difference between the PCR estimates of M. bovis in both blood and milk in dairy cattle, where the sensitivity of PCR conducted on blood and milk samples was 53–95% and 1–60% while its specificity was 95–100% and 43–99% for each sample, respectively ^[36]. The authors attributed these differences to the type of sampling, that largely affect the PCR sensitivity and specificity estimates. However, further studies about bacteremia and the time of dissemination of M. bovis in blood stream is needed to detect the proper sampling time because most of TST and IGRA reactor animals failed to be detected by PCR in blood samples ^[81]. Despite the high risk of disseminated infection of M. bovis, bacteremia has been assumed to be rare in cattle ^[82].

In the light of the aforementioned, bTB lacks a definitive gold standard diagnostic test ^[44] since no single diagnostic method is able to detect all infected animals ^[78]. In addition, several classical tests based on growth, phenotypic, and biochemical properties had been assayed to discriminate between members of MBTC ^[9]. However, these tests were inaccurate, slow, time consuming, cumbersome, and cannot be performed in any laboratory ^[39]. Hence, the usage of advanced molecular diagnostic methods has been highlighted in recent years, especially because all members of MBTC share about 99.9% of nucleotide identity ^[83]. Spoligotyping and MIRU-VNTR profiling have been widely used as molecular typing methods of MBTC members ^[84]^[85]. However, due to a low discriminatory power, these methods have limitations for phylogenetic studies so, the advances in the high-throughput sequencing technology become urgent over the last decade ^[86].

Whole Genome Sequencing (WGS) provides new insights into dynamics of disease transmission, host-pathogen interactions, and comparative analysis that elucidates key differences between the animal and human mycobacteria ^[39]^[87]. Abdelaal et al. performed WGS on Egyptian M. bovis isolates from Nile Delta and reported a predominance of isolates which were closely related to clonal complex (European 2).

Clarke et al. investigated an in-field sampling technique for rapid, safe detection of M. bovis in buffalo tissues ^[88]. The authors recommended the use of PrimeStore^® Molecular Transport Medium, in combination with Xpert^® MTB/RIF Ultra assay (an automated cartridge-based semi-quantitative nested RT-PCR assay that detects DNA of MBTC and rifampicin (RIF) resistance in clinical specimens) as a safe and rapid PM screening test for M. bovis in buffaloes. In humans, Xpert MTB/RIF has recently become a significant breakthrough in TB diagnosis ^[89]. This test can detect MBTC DNA and mutations associated with antitubercular drug resistance.

Kapalamula et al. developed a loop-mediated isothermal amplification (LAMP) method for specific identification of M. bovis. This LAMP method was able to detect M. bovis within 40 min following incubation and results could be read with the naked eye following development of a color change ^[90]. .

Additional tools for diagnosing TB, such as lymph node biopsy and tracheobronchial aspirates, have been successfully implemented in wildlife ruminants. Didkowska et al. used tracheobronchial aspirates and ultrasound-guided biopsies for diagnosis of TB in European bison ^[91]. In addition, Didkowska et al. used endoscopy (bronchoscopy) as an additional tool for diagnosing tuberculosis in European bison, especially in highly valuable animals, and to assess the stage of the disease ^[92]. In human, sputum samples are widely used for diagnosing pulmonary tuberculosis ^[93].

References

Vordermeier, H.M.; Jones, G.J.; Buddle, B.M.; Hewinson, R.G.; Villarreal-Ramos, B. Bovine tuberculosis in cattle: Vaccines, DIVA tests, and host biomarker discovery. Annu. Rev. Anim. Biosci. 2016, 4, 87–109.
Radostits, O.M.; Gay, C.C.; Blood, D.C.; Hinchliff, K.W. Veterinary Medicine. A Textbook of the Diseases of Cattle, Sheep, Goats and Horses, 8th ed.; Ballier Tindals: London, UK, 2007; pp. 830–838.
Klepp, L.I.; Eirin, M.E.; Garbaccio, S.; Soria, M.; Bigi, F.; Blanco, F.C. Identification of bovine tuberculosis biomarkers to detect tuberculin skin test and IFNγ release assay false negative cattle. Res. Vet. Sci. 2019, 122, 7–14.
Abdellrazeq, G.S.; Elnaggar, M.M.; Osman, H.S.; Davis, W.C.; Singh, M. Prevalence of Bovine Tuberculosis in Egyptian Cattle and the Standardization of the Interferon-gamma Assay as an Ancillary Test. Transbound. Emerg. Dis. 2016, 63, 497–507.
Stear, M. OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals (Mammals, Birds and Bees) 5th Edn. Volumes 1 and 2. World Organization for Animal Health 2004. ISBN 92 9044 622 6.€ 140. Parasitology 2005, 130, 727.
Elnaggar, M.M.; Abdellrazeq, G.S.; Elsisy, A.; Mahmoud, A.H.; Shyboub, A.; Sester, M.; Davis, W.C. Evaluation of antigen specific interleukin-1β as a biomarker to detect cattle infected with Mycobacterium bovis. Tuberculosis 2017, 105, 53–59.
Broughan, J.M.; Judge, J.; Ely, E.; Delahay, R.J.; Wilson, G.; Clifton-Hadley, R.S.; Goodchild, A.V.; Bishop, H.; Parry, J.E.; Downs, S.H. A review of risk factors for bovine tuberculosis infection in cattle in the UK and Ireland. Epidemiol. Infect. 2016, 144, 2899–2926.
Constable, P.D.; Hinchcliff, K.W.; Done, S.H.; Grünberg, W. Veterinary Medicine-e-Book: A Textbook of the Diseases of Cattle, Horses, Sheep, Pigs and Goats; Elsevier: Amsterdam, The Netherlands, 2016.
De la Rua-Domenech, R.; Goodchild, A.T.; Vordermeier, H.M.; Hewinson, R.G.; Christiansen, K.H.; Clifton-Hadley, R.S. Ante mortem diagnosis of tuberculosis in cattle: A review of the tuberculin tests, γ-interferon assay and other ancillary diagnostic techniques. Res. Vet. Sci. 2006, 81, 190–210.
Picasso-Risso, C.; Grau, A.; Bakker, D.; Nacar, J.; Mínguez, O.; Perez, A.; Alvarez, J. Association between results of diagnostic tests for bovine tuberculosis and Johne’s disease in cattle. Vet. Rec. 2019, 185, 693.
Byrne, A.W.; Graham, J.; Brown, C.; Donaghy, A.; Guelbenzu-Gonzalo, M.; McNair, J.; McDowell, S.W. Modelling the variation in skin-test tuberculin reactions, post-mortem lesion counts and case pathology in tuberculosis-exposed cattle: Effects of animal characteristics, histories and co-infection. Transbound. Emerg. Dis. 2018, 65, 844–858.
Howell, A.K.; McCann, C.M.; Wickstead, F.; Williams, D.J. Co-infection of cattle with Fasciola hepatica or F. gigantica and Mycobacterium bovis: A systematic review. PLoS ONE 2019, 14, e0226300.
Borham, M.; Oreiby, A.; El-Gedawy, A.; Hegazy, Y.; Al-Gaabary, M. Tuberculin test errors and its effect on detection of bovine tuberculosis. J. Hell. Vet. Med. 2021, 72, 3263–3270.
Bezos, J.; Casal, C.; Romero, B.; Schroeder, B.; Hardegger, R.; Raeber, A.J.; Domínguez, L. Current ante-mortem techniques for diagnosis of bovine tuberculosis. Res. Vet. Sci. 2014, 97, S44–S52.
Parlane, N.A.; Chen, S.; Jones, G.J.; Vordermeier, H.M.; Wedlock, D.N.; Rehm, B.H.; Buddle, B.M. Display of antigens on polyester inclusions lowers the antigen concentration required for a bovine tuberculosis skin test. Clin. Vaccine Immunol. 2016, 23, 19–26.
Picasso-Risso, C.; Alvarez, J.; VanderWaal, K.; Kinsley, A.; Gil, A.; Wells, S.J.; Perez, A. Modelling the effect of test-and-slaughter strategies to control bovine tuberculosis in endemic high prevalence herds. Transbound. Emerg. Dis. 2021, 68, 1205–1215.
Didkowska, A.; Orłowska, B.; Krajewska-Wędzina, M.; Krzysiak, M.; Bruczyńska, M.; Wiśniewski, J.; Anusz, K. Intra-palpebral tuberculin skin test and interferon gamma release assay in diagnosing tuberculosis due to Mycobacterium caprae in European Bison (Bison bonasus). Pathogens 2022, 11, 260.
Pollock, J.M.; Neill, S.D. Mycobacterium bovis infection and tuberculosis in cattle. Vet. J. 2002, 163, 115–127.
El-Sawalhy, A. Veterinary Infectious Diseases in Domestic Animals, 3rd ed.; Vetbook: Cairo, Egypt, 2012; pp. 305–308.
Belinda, S.T.; Erin, L.G. Miscellaneous Infectious Diseases. In Rebhun’s Diseases of Dairy Cattle, 3rd ed.; Elsevier: Amsterdam, The Netherlands, 2018; pp. 745–746.
Pascual-Linaza, A.V.; Gordon, A.W.; Stringer, L.A.; Menzies, F.D. Efficiency of slaughterhouse surveillance for the detection of bovine tuberculosis in cattle in Northern Ireland. Epidemiol. Infect. 2017, 145, 995–1005.
Abbate, J.M.; Arfuso, F.; Iaria, C.; Arestia, G.; Lanteri, G. Prevalence of bovine tuberculosis in slaughtered cattle in Sicily, Southern Italy. Animals 2020, 10, 1473.
Domingo, M.; Vidal, E.; Marco, A. Pathology of bovine tuberculosis. Res. Vet. Sci. 2014, 97, S20–S29.
Aboukhassib, H.; Haraji, M.; Bouslikhane, M.; Bitar, A. Bovine tuberculosis: Clinical presentation and diagnosis. J. Bacteriol. Mycol. Open Access 2016, 3, 214–217.
Ahmad, I.; Kudi, C.A.; Abdulkadir, A.I.; Saidu, S.N.A. Occurrence and distribution of bovine TB pathology by age, sex, and breed of cattle slaughtered in Gusau Abattoir, Zamfara State Nigeria. Trop. Anim. Health Prod. 2017, 49, 583–589.
Kuria, J.K. Diseases Caused by Bacteria in Cattle: Tuberculosis. In Bacterial Cattle Diseases; IntechOpen: London, UK, 2019.
Genzebu, A.T.; Menghistu, H.T.; Weldearegay, Y.H. A case of miliary tuberculosis in a Holestein Fresian dairy cow, Mekelle, Tigray, Ethiopia. Ethiop. Vet. J. 2018, 22, 121–129.
Oreiby, A.F.; Osman, S.A.; Hegazy, Y.; Ghanem, Y.A.; Al-Gaabary, M.H. Caseous lymphadenitis in small ruminants: Descriptive, epidemiological and clinical studies. Kafrelsheikh Vet. Med. J. 2013, 11, 41–61.
Borham, M.A.; Oreiby, A.F.; El-Gedawy, A.A.; Al-Gaabary, M.H. Serological Surveillance of Caseous Lymphadenitis in Sudanese and Somali Camels Slaughtered at Al-warraq Abattoir, Giza, Egypt. World’s Vet. J. 2016, 6, 89–94.
Borham, M.; Oreiby, A.; El-Gedawy, A.; Al-Gaabary, M. Caseous Lymphadenitis in Sudanese and Somalian Camels Imported for Meat Consumption in Egypt. Alex. J. Vet. Sci. 2017, 55, 52–59.
Sinclair, J.A.; Dawson, K.L.; Buddle, B.M. The effectiveness of parallel gamma-interferon testing in New Zealand’s bovine tuberculosis eradication programme. Prev. Vet. Med. 2016, 127, 94–99.
Algammal, A.M.; Wahdan, A.; Elhaig, M.M. Potential efficiency of conventional and advanced approaches used to detect Mycobacterium bovis in cattle. Microb. Pathog. 2019, 134, 103574.
Buddle, B.M.; De Lisle, G.W.; Pfeffer, A.; Aldwell, F.E. Immunological responses and protection against Mycobacterium bovis in calves vaccinated with a low dose of BCG. Vaccine 1995, 13, 1123–1130.
Oreiby, A.F.; Hegazy, Y.M. Diagnosis of ovine caseous lymphadenitis by blood and milk gamma interferon assays. Small Rumin. Res. 2016, 144, 109–112.
Ryan, T.J.; Buddle, B.M.; De Lisle, G.W. An evaluation of the gamma interferon test for detecting bovine tuberculosis in cattle 8 to 28 days after tuberculin skin testing. Res. Vet. Sci. 2000, 69, 57–61.
Elsohaby, I.; Mahmmod, Y.S.; Mweu, M.M.; Ahmed, H.A.; El-Diasty, M.M.; Elgedawy, A.A.; El Hofy, F.I. Accuracy of PCR, mycobacterial culture and interferon-γ assays for detection of Mycobacterium bovis in blood and milk samples from Egyptian dairy cows using Bayesian modelling. Prev. Vet. Med. 2020, 181, 105054.
Clarke, C.; Cooper, D.; Goosen, W.J.; McFadyen, R.; Warren, R.M.; van Helden, P.D.; Miller, M.A. Antigen-specific interferon-gamma release is decreased following the single intradermal comparative cervical skin test in African buffaloes (Syncerus caffer). Vet. Immunol. Immunopathol. 2018, 201, 12–15.
Bernitz, N.; Goosen, W.J.; Clarke, C.; Kerr, T.J.; Higgitt, R.; Roos, E.O.; Miller, M.A. Parallel testing increases detection of Mycobacterium bovis-infected African buffaloes (Syncerus caffer). Vet. Immunol. Immunopathol. 2018, 204, 40–43.
Ramos, D.F.; Silva, P.E.A.; Dellagostin, O.A. Diagnosis of bovine tuberculosis: Review of main techniques. Braz J. Biol. 2015, 75, 830–837.
Alvarez, A.H.; Gutiérrez-Ortega, A.; Gómez-Entzin, V.; Pérez-Mayorga, G.; Naranjo-Bastién, J.; González-Martínez, V.; Hinojoza-Loza, E. Assessment of antigenic supplementation of bovine purified protein derivative for diagnosis of subclinical infection with Mycobacterium bovis in cattle. Microb. Pathog. 2017, 108, 114–121.
Palmer, M.V.; Thacker, T.C.; Rabideau, M.M.; Jones, G.J.; Kanipe, C.; Vordermeier, H.M.; Waters, W.R. Biomarkers of cell-mediated immunity to bovine tuberculosis. Vet. Immunol. Immunopathol. 2020, 220, 109988.
Vordermeier, H.M.; Cockle, P.J.; Whelan, A.O.; Rhodes, S.; Hewinson, R.G. Toward the development of diagnostic assays to discriminate between Mycobacterium bovis infection and bacille Calmette-Guerin vaccination in cattle. Clin. Infect. Dis. 2000, 30, S291–S298.
Fontana, S.; Pacciarini, M.; Boifava, M.; Pellesi, R.; Casto, B.; Gastaldelli, M.; Boniotti, M.B. Development and evaluation of two multi-antigen serological assays for the diagnosis of bovine tuberculosis in cattle. J. Microbiol. Methods 2018, 153, 118–126.
Soares Filho, P.M.; Ramalho, A.K.; de Moura Silva, A.; Hodon, M.A.; de Azevedo Issa, M.; Júnior, A.A.F.; Leite, R.C. Evaluation of post-mortem diagnostic tests’ sensitivity and specificity for bovine tuberculosis using Bayesian latent class analysis. Res. Vet. Sci. 2019, 125, 14–23.
Casal, C.; Díez-Guerrier, A.; Álvarez, J.; Rodriguez-Campos, S.; Mateos, A.; Linscott, R.; O’Brien, A. Strategic use of serology for the diagnosis of bovine tuberculosis after intradermal skin testing. Vet. Microbiol. 2014, 170, 342–351.
Van der Heijden, E.M.; Cooper, D.V.; Rutten, V.P.; Michel, A.L. Mycobacterium bovis prevalence affects the performance of a commercial serological assay for bovine tuberculosis in African buffaloes. Comp. Immunol. Microbiol. Infect. Dis. 2020, 70, 101369.
Nunez-Garcia, J.; Downs, S.H.; Parry, J.E.; Abernethy, D.A.; Broughan, J.M.; Cameron, A.R.; More, S.J. Meta-analyses of the sensitivity and specificity of ante-mortem and post-mortem diagnostic tests for bovine tuberculosis in the UK and Ireland. Prev. Vet. Med. 2018, 153, 94–107.
Souza, I.I.F.; de Arruda Rodrigues, R.; Jorge, K.S.G.; Silva, M.R.; Lilenbaum, W.; Vidal, C.E.S.; Araujo, F.R. ELISA using a recombinant chimera of ESAT-6/MPB70/MPB83 for Mycobacterium bovis diagnosis in naturally infected cattle. J. Vet. Med. Sci. 2019, 81, 9–14.
Griffa, N.; Moyano, R.D.; Canal, A.M.; Travería, G.E.; Santangelo, M.P.; Alonso, N.; Romano, M.I. Development and diagnostic validation of an ELISA based on an antigenic mixture for the detection of bovine tuberculosis. Vet. J. 2020, 256, 105426.
Casal, C.; Infantes, J.A.; Risalde, M.A.; Díez-Guerrier, A.; Domínguez, M.; Moreno, I.; Gortázar, C. Antibody detection tests improve the sensitivity of tuberculosis diagnosis in cattle. Res. Vet. Sci. 2017, 112, 214–221.
McCallan, L.; Brooks, C.; Barry, C.; Couzens, C.; Young, F.J.; McNair, J.; Byrne, A.W. Serological test performance for bovine tuberculosis in cattle from herds with evidence of on-going infection in Northern Ireland. PLoS ONE 2021, 16, e0245655.
Waters, W.R.; Palmer, M.V.; Stafne, M.R.; Bass, K.E.; Maggioli, M.F.; Thacker, T.C.; Greenwald, R. Effects of serial skin testing with purified protein derivative on the level and quality of antibodies to complex and defined antigens in Mycobacterium bovis-infected cattle. Clin. Vaccine Immunol. 2015, 22, 641–649.
Al-Fattli, H.H.H. The clinical and serological diagnosis of Mycobacterium bovis in blood and milk serums of lactating cows by IDEXX ELISA test in Wasit and Dhi-Qar provinces/Iraq. J. Contemp. Med. Sci. 2016, 2, 70–73.
Trost, B.; Stuber, T.; Surujballi, O.; Nelson, J.; Robbe-Austerman, S.; Smith, N.H.; Griebel, P. Investigation of the cause of geographic disparities in IDEXX ELISA sensitivity in serum samples from Mycobacterium bovis-infected cattle. Sci. Rep. 2016, 6, 22763.
Elsohaby, I.; Ahmed, H.A.; El-Diasty, M.M.; Elgedawy, A.A.; Mahrous, E.; El Hofy, F.I. Serological and molecular evidence of Mycobacterium bovis in dairy cattle and dairy farm workers under the intensive dairy production system in Egypt. J. Appl. Microbiol. 2020, 129, 1207–1219.
Gormley, E.; Corner, L.A.L.; Costello, E.; Rodriguez-Campos, S. Bacteriological diagnosis and molecular strain typing of Mycobacterium bovis and Mycobacterium caprae. Res. Vet. Sci. 2014, 97, S30–S43.
Fauci, A.S.; Braunwald, E.; Kasper, D.L.; Hauser, S.; Longo, D.; Jameson, J. Harrisons Principles of Internal Medicine; McGraw-Hil: New York, NY, USA, 2008; pp. 1543–1571.
Anne, N.S.; Ronald, B.S.M.; Kumar, T.S.; Kannan, P.; Thangavelu, A. Molecular identification of Mycobacterium tuberculosis in cattle. Vet. Microbiol. 2017, 198, 81–87.
Worku, A.; Abreham, S.; Hailu, M.; Mamo, G.; Ameni, G.; Tsegaye, S. Cross-sectional study and comparison of different diagnostic methods of bovine tuberculosis in Gondar Elfora abattoir, Ethiopia. Mycobact. Dis. 2016, 6, 1068–2161.
Mohamed, M.; Moussa, L.M.; Mohamed, K.F.; Samir, A.; Nasr, E.A.; Ashgan, M.H.; Hatem, M.E. BACTEC MGIT 960 TM system for screening of Mycobacterium tuberculosis complex among cattle. Afr. J. Biotechnol. 2011, 10, 13919–13923.
Corner, L.A.L.; Gormley, E.; Pfeiffer, D.U. Primary isolation of Mycobacterium bovis from bovine tissues: Conditions for maximising the number of positive cultures. Vet. Microbiol. 2012, 156, 162–171.
Issa, M.D.A.; Soares Filho, P.M.; Fonseca Júnior, A.A.; Hodon, M.A.; Santos, L.C.D.; Reis, J.K.P.D.; Cerqueira Leite, R. Comparative study of Mycobacterium bovis primary isolation methods. Braz. J. Microbiol. 2017, 48, 139–144.
Albernaz, T.T.; Oliveira, C.M.C.; da Silva Lima, D.H.; e Silva, N.D.S.; Cardoso, D.P.; Lopes, C.T.A.; Barbosa, J.D. Comparison of the tuberculin test, histopathological examination, and bacterial culture for the diagnosis of tuberculosis (Mycobacterium bovis) in buffaloes (Bubalus bubalis) in Brazil. Trop. Anim. Health Prod. 2015, 47, 1153–1159.
Mayer, F.Q.; Reis, E.M.D.; Bezerra, A.V.A.; Rodrigues, R.O.; Michel, T.; Cerva, C.; Bertagnolli, A.C. Nasal swab real-time PCR is not suitable for in vivo diagnosis of bovine tuberculosis. Pesqui. Vet. Bras. 2017, 37, 549–554.
Canal, A.M.; Pezzone, N.; Cataldi, A.; Zumarraga, M.; Larzabal, M.; Garbaccio, S.; Rodriguez-Bertos, A. Immunohistochemical detection of pro-inflammatory and anti-inflammatory cytokines in granulomas in cattle with natural Mycobacterium bovis infection. Res. Vet. Sci. 2017, 110, 34–39.
McKinley, T.J.; Lipschutz-Powell, D.; Mitchell, A.P.; Wood, J.L.; Conlan, A.J. Risk factors and variations in detection of new bovine tuberculosis breakdowns via slaughterhouse surveillance in Great Britain. PLoS ONE 2018, 13, e0198760.
Michelet, L.; de Cruz, K.; Karoui, C.; Tambosco, J.; Moyen, J.L.; Hénault, S.; Boschiroli, M.L. Second line molecular diagnosis for bovine tuberculosis to improve diagnostic schemes. PLoS ONE 2018, 13, e0207614.
Brahma, D.; Narang, D.; Chandra, M.; Gupta, K.; Singh, A.; Kaur, G. Diagnosis of Mycobacterial infections (Tuberculosis and Paratuberculosis) in tissue samples using molecular (in-house multiplex PCR, PCR and TaqMan real-time PCR), histopathology and immunohistochemical techniques. Trop. Biomed. 2017, 34, 911–927.
Elsayed, M.S.A.E.; Amer, A. The rapid detection and differentiation of Mycobacterium tuberculosis complex members from cattle and water buffaloes in the delta area of Egypt, using a combination of real-time and conventional PCR. Mol. Biol. Rep. 2019, 46, 3909–3919.
Warren, R.M.; Gey van Pittius, N.C.; Barnard, M.; Hesseling, A.; Engelke, E.; De Kock, M.; Van Helden, P.D. Differentiation of Mycobacterium tuberculosis complex by PCR amplification of genomic regions of difference. Int. J. Tuberc. Lung Dis. 2006, 10, 818–822.
Thacker, T.C.; Harris, B.; Palmer, M.V.; Waters, W.R. Improved specificity for detection of Mycobacterium bovis in fresh tissues using IS 6110 real-time PCR. BMC Vet. Res. 2011, 7, 50.
Rakotosamimanana, N.; Rabodoarivelo, M.S.; Palomino, J.C.; Martin, A.; Razanamparany, V.R. Exploring tuberculosis by molecular tests on DNA isolated from smear microscopy slides. Int. J. Infect. Dis. 2017, 56, 248–252.
Kumar, M.; Sharma, S.; Ram, A.B.; Khan, I.A. Efficient mycobacterial DNA extraction from clinical samples for early diagnosis of tuberculosis. Int. J. Tuberc. Lung Dis. 2010, 14, 847–851.
Ikuta, C.Y.; Oliveira, D.C.R.; de Souza, G.O.; de Souza Filho, A.F.; Grisi-Filho, J.H.H.; Heinemann, M.B.; Neto, J.S.F. Comparison of DNA extraction protocols to detect Mycobacterium bovis in bovine tissue by PCR. Semin. Cienc. Agrar. 2016, 37, 3709–3718.
Cardoso, M.A.; Cardoso, R.F.; Hirata, R.D.C.; Hirata, M.H.; Leite, C.Q.F.; Santos, A.C.B.; Lonardoni, M.V.C. Direct detection of Mycobacterium bovis in bovine lymph nodes by PCR. Zoonoses Public Health 2009, 56, 465–470.
Yoshikawa, H.; Dogruman-Ai, F.; Turk, S.; Kustimur, S.; Balaban, N.; Sultan, N. Evaluation of DNA extraction kits for molecular diagnosis of human Blastocystis subtypes from fecal samples. Parasitol. Res. 2011, 109, 1045–1050.
Moura, A.; Hodon, M.A.; Soares Filho, P.M.; Issa, M.D.A.; Oliveira, A.P.F.D.; Fonseca Júnior, A.A. Comparison of nine DNA extraction methods for the diagnosis of bovine tuberculosis by real time PCR. Ciênc. Rural 2016, 46, 1223–1228.
Carvalho, R.C.T.; Furlanetto, L.V.; Maruyama, F.H.; de Araújo, C.P.; Barros, S.L.B.; do Nascimento Ramos, C.A.; de Souza Figueiredo, E.E. Evaluation of the efficiency of nested q-PCR in the detection of Mycobacterium tuberculosis complex directly from tuberculosis-suspected lesions in post-mortem macroscopic inspections of bovine carcasses slaughtered in the state of Mato Grosso, Brazil. Meat Sci. 2015, 106, 11–15.
Helmy, N.M.; Abdel-Moghney, A.R.F.; Atia, M.A. Evaluation of Different PCR-Based Techniques in Diagnosis of Bovine Tuberculosis in Infected Cattle Lymph Nodes. Am. J. Microbiol. Biotechnol. 2015, 2, 75–81.
Mataragka, A.; Fytani, V.; Ikonomopoulos, J.; Soptirakoglou, K.; Katsiolis, A.; Dile, C. Assessment of the Use of PCR as an Early Diagnostic Indicator of Bovine Tuberculosis in Dairy Farms. Mycobact. Dis. 2019, 9, 1068–2161.
Brahma, D.; Narang, D.; Chandra, M.; Filia, G.; Singh, A.; Singh, S.T. Diagnosis of Bovine Tuberculosis by Comparative Intradermal Tuberculin Test, Interferon Gamma Assay and esxB (CFP-10) PCR in Blood and Lymph Node Aspirates. Open J. Vet. Med. 2019, 9, 55–65.
Maggioli, M.F. A bloody evidence: Is Mycobacterium bovis bacteraemia frequent in cattle? Virulence 2016, 7, 748–750.
Villarreal-Ramos, B.; Berg, S.; Whelan, A.; Holbert, S.; Carreras, F.; Salguero, F.J.; Smyth, A. Experimental infection of cattle with Mycobacterium tuberculosis isolates shows the attenuation of the human tubercle bacillus for cattle. Sci. Rep. 2018, 8, 894.
Carvalho, R.C.T.; Vasconcellos, S.E.G.; de Azevedo Issa, M.; Soares Filho, P.M.; Mota, P.M.P.C.; de Araujo, F.R.; Paschoalin, V.M.F. Molecular typing of Mycobacterium Bovis from cattle reared in Midwest Brazil. PLoS ONE 2016, 11, e0162459.
Ghebremariam, M.K.; Hlokwe, T.; Rutten, V.P.; Allepuz, A.; Cadmus, S.; Muwonge, A.; Michel, A.L. Genetic profiling of Mycobacterium bovis strains from slaughtered cattle in Eritrea. PLoS Negl. Trop. Dis. 2018, 12, e0006406.
Da Conceição, M.L.; Conceição, E.C.; Furlaneto, I.P.; da Silva, S.P.; dos Santos Guimarães, A.E.; Gomes, P.; da Costa Francez, L. Phylogenomic perspective on a unique Mycobacterium bovis clade dominating bovine tuberculosis infections among cattle and buffalos in Northern Brazil. Sci. Rep. 2020, 10, 1747.
Abdelaal, H.F.; Spalink, D.; Amer, A.; Steinberg, H.; Hashish, E.A.; Nasr, E.A.; Talaat, A.M. Genomic polymorphism associated with the emergence of virulent isolates of Mycobacterium bovis in the Nile Delta. Sci. Rep. 2019, 9, 11657.
Clarke, C.; Smith, K.; Goldswain, S.J.; Helm, C.; Cooper, D.V.; Kerr, T.J.; Goosen, W.J. Use of PrimeStore® Molecular Transport Medium and Xpert MTB/RIF Ultra for Rapid Detection of Mycobacterium Bovis in African buffaloes (Syncerus Caffer); Research Square: Durham, NC, USA, 2021; pp. 1–11.
Pillay, S.; Steingart, K.R.; Davies, G.R.; Chaplin, M.; De Vos, M.; Schumacher, S.G.; Theron, G. Xpert MTB/XDR for detection of pulmonary tuberculosis and resistance to isoniazid, fluoroquinolones, ethionamide, and amikacin. Cochrane Database Syst. Rev. 2022, 5, CD014841.
Kapalamula, T.F.; Thapa, J.; Akapelwa, M.L.; Hayashida, K.; Gordon, S.V.; Hang’Ombe, B.M.; Nakajima, C. Development of a loop-mediated isothermal amplification (LAMP) method for specific detection of Mycobacterium bovis. PLoS Negl. Trop. Dis. 2021, 15, e0008996.
Didkowska, A.; Orłowska, B.; Witkowski, L.; Olbrych, K.; Brzezińska, S.; Augustynowicz-Kopeć, E.; Anusz, K. Biopsy and tracheobronchial aspirates as additional tools for the diagnosis of bovine tuberculosis in living European bison (Bison bonasus). Animals 2020, 10, 2017.
Didkowska, A.; Dziekan, P.; Czujkowska, A.; Bereznowski, A.; Witkowski, L.; Orłowska, B.; Anusz, K. The first visually-guided bronchoscopy in European bison (Bison bonasus)–An additional tool in the diagnosis of bovine tuberculosis? Vet. Anim. Sci. 2021, 12, 100174.
Mulengwa, D.L.; Monyama, M.C.; Lebelo, S.L. Evaluation of the GeneXpert MTB/RIF assay performance in sputum samples with various characteristics from presumed pulmonary tuberculosis patients in Shiselweni region, Eswatini. Inf. Dis. 2022, 54, 170–177.

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