| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
 |
ABSTRACT |
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
|
INTRODUCTION |
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Cattle become infected mainly by respiratory route and remain asymptomatic during the first few months after infection, but symptoms can appear when the delicate balance between the host and the infectious agent is lost because of stress factors such as immunosuppression or malnutrition.1 After infection, nodular granulomas, known as tubercles, start to develop; these can occur in any tissue but are most frequently observed in lymph nodes, lungs, intestines, liver, spleen, pleura, and peritoneum. Clinical evidence of TB is usually lacking, and symptoms, if present, are not specific, e.g., sub-febrile temperatures, coughing, fatigue, loss of appetite, and reduced milk production.4
The status of BTB in Ecuador is not documented or clearly quantified because of several factors. These include lack of proper recording of positive cases, limited use of diagnostic tests, and insufficient veterinary inspection in most of the slaughterhouses. Isolated surveys carried out on BTB in cattle report a prevalence of 3.91% in the northern part of the country (Andino-Ashqui, unpublished data). In 2001, 46.55 cases of human TB per 100,000 inhabitants were reported by the Ministry of Public Health.5 Nevertheless, studies have not been conducted to quantify cases of human TB caused by M. bovis. In developing countries, the frequency and involvement of M. bovis in non-pulmonary TB is largely unknown because of limited laboratory facilities for culture and identification of tubercle bacilli.6
The standard method used for routine diagnosis of BTB is the tuberculin test,7 which consists of reading the skin reaction to an intradermally injected purified protein derivative (PPD). Differentiation of the causative Mycobacterium species usually requires in vitro growth of the organism. Alternatively, the polymerase chain reaction (PCR) has been used with considerable success,8 and has significantly reduced the time needed to confirm diagnosis in suspected cases. The purpose of this study was to evaluate the situation of BTB in Mejia canton, a major dairy cattle production region in Ecuador.
|
MATERIALS AND METHODS |
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Tuberculin skin test.
A total of 1,012 randomly selected cross-bred dairy cattle improved by Holstein-Friesian cross-breeding from 59 farms were tested by an intradermal tuberculin skin test. Based on their herd size, dairy farms were grouped as large (more than 70 cattle), medium (25–70 cattle), and small (1–25 cattle). Tuberculin tests were restricted to animals more than six months of age, i.e., 22 animals randomly selected from each large or medium farm and all animals from small farms. All animals were ear-tagged. Injection sites were cleaned and disinfected; swellings were measured with callipers immediately after inoculation and 72 hours later. A negative result in the SST was a swelling 
3 mm, a suspicious result was a swelling between 3 and 5 mm, and a positive result was a swelling > 5 mm.
To confirm positive or suspicious STT reactions, a comparative intradermal tuberculin test with bovine and avian tuberculin (CTT) was carried out after 60 days. The interpretation of the CTT after 72 hours is based on the comparison between the subsequent swellings. Reactions to bovine tuberculin 5 mm and > 4 mm larger than the reaction to avian tuberculin were considered positive.
For the STT, 0.1 of mL bovine tuberculin (PPD-protein M. bovis, strain AN-5 at 25,000 IU/mL; CZ Veterinaria SA, Porriño Spain) was injected intradermally 7 cm from the base of the ventral side of the tail. The CTT injections were administered intradermally on the left side of the neck with the same bovine tuberculin and 0.1 mL of avian PPD tuberculin (0.5 mg/mL, 25,000 IU/mL; Servicio Nacional de Sanidad, Buenos Aires, Argentina) in previously shaven injection sites 12 cm apart.
Isolation of Mycobacterium spp. One hundred twenty-five biopsy specimens (lymph nodes, lungs, kidneys, and mammary glands) were taken from 40 cattle in the local district slaughterhouse with suspicious macroscopic lesions. In addition, 11 and 18 necropsy samples, respectively, were obtained from two cattle that were slaughtered (no. 65 and no. 1555) and positive in the CTT.
All specimens were stored in 2-mL Eppendorf (Hamburg, Germany) tubes containing Dubos broth (no. 0385-17-6; Difco Laboratories, Detroit, MI) supplemented with PANTA (Bactec PANTA Plus Kit no. 440 476 4; Becton Dickinson, Franklin Lakes, NJ) to preserve the mycobacteria and retard growth of contaminants.9 Samples were transported at 5°C and stored at –20°C at the Department of Microbiology, Unit of Mycobacteriology, Institute for Tropical Medicine in Antwerp until processed.
In vitro culture and phenotypic identification. All samples were decontaminated using the inverted Petroff method, i.e., 20-minute decontamination with 1 N HCl followed by alkaline neutralization (1 N NaOH).10 Subsequently, Ziehl-Neelsen staining and inoculation into Löwenstein-Jensen and Stonebrink11 media was conducted. In vitro identification of mycobacteria was carried out using biochemical (i.e., niacin production, nitrate reduction, urease production) and inhibition tests (thiophene-2-carboxylic hydrazide) and pigmentation and colony morphology as described by Lévy-Frébault and Portaels.12
Polymerase chain reaction. Necropsy samples from the two CTT-positive cattle were examined by PCR. A nested PCR was performed as described by Portaels and others13 with primers for the 16S ribosomal RNA gene.14 Primers P1: 5'-TGCTTAACACATGCAAGTCG-3' and P2: 5'-TGAGAT TTCACGAACAACGC-3' were used at 56°C in the first PCR as general primers for identifying the Mycobacterium genus. Primers P3: 5'-AACCCGGACCTTCGTCGATG-3' and P9: 5'-CATGTCTTGTGGTGGAAAGCGC-3' were used at 66°C in the second PCR for specific identification of the M. tuberculosis complex. The final PCR product was approximately 540 basepairs.
DNA was extracted from tissues as reported by Portaels and others9 using proteinase K. Briefly, 50 µL of proteinase K (no. V3021, 20 mg/mL; Promega, Madison, WI) and 250 µL of lysis buffer (1.6 M guanidine hydrochloride, 60 mM Tris, pH 7.4, 1% Triton, 60 mM EDTA, 10% Tween-20) were added to the decontaminated samples (250 µL) and incubated for one hour at 60°C with shaking (200 rpm). Approximately 80 mg of glass beads was then added, samples were sonicated for five minutes at room temperature in a water bath sonicator (14 kHz; Branson 1200; Branson Ultrasonics Corporation, Danbury, CT), 40 µL of acidified diatomaceous earth solution was added, and the suspensions were incubated for a maximum of two hours at 37°C with shaking (200 rpm). The suspensions were extracted twice with 70% ethanol and once with acetone with intermediate short centrifugations. The pellets were dried at 50°C in a heating block (Dri-Bath 16500; Thermolyne, Merck-Belgolabo, Belgium) and resuspended in 90 µL of TE buffer (10 mM Tris, 1 mM EDTA, pH 8), followed by incubation for 20 minutes at 55°C while shaking to obtain complete homogenization. The sample was briefly centrifuged and 50 µL of supernatant was transferred to a new tube.
For the first PCR, 10 µL of the DNA extract was added to 40 µL of PCR mixture containing 50 pmol of each primer, 1 unit of Ampli Taq DNA polymerase (no. M1668; Promega), 200 µM dNTP (no. 27-2094-02; Pharmacia, Uppsala, Sweden), 25 µL of buffer, and 7 µL of milli-Q water (Millipore, Billerica, MA) and overlaid with two drops of mineral oil (no. 50138; ICN, Costa Mesa, CA). A negative control (milli-Q water) and positive control (DNA at a specific concentration) were included in each PCR. Denaturation was at 94°C for 5 minutes, amplification for 40 cycles at 94°C for 45 seconds, 56°C for 45 seconds, and 72°C for 45 seconds, and final extension at 72°C for 10 minutes.
For the second PCR, 1 µL of the first PCR product was amplified in a 25-µL reaction mixture containing 25 pmol of each primer, 0.5 units of Ampli Taq DNA polymerase, 200 µM dNTP, 12.5 µL of buffer, 8.5 µL of milli-Q water and overlaid with one drop of mineral oil. Amplification was composed of 25 cycles as described for the first PCR with an annealing temperature of 66°C. A total of 75 µL of amplified DNA plus 2.5 µL of loading buffer (Fermentas, St. Leon-Rot, Germany) and a molecular size marker were subjected to electrophoresis on a 2% agarose gel (no. EP-0010-10; Eurogentec, Seraing, Belgium) in 0.5% TAE buffer (1 mM EDTA, pH 8.0, 40 mM Tris-acetate). Bands were detected by staining with ethidium bromide (Bio-Rad Laboratories, Hercules, CA) and transillumination with ultraviolet light. The pattern of bands obtained was compared with those of negative and positive controls.
|
RESULTS |
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
). To confirm these results or discard possible false-positive results, the CTT was used 60 days after the STT and was conducted only with positive and suspected positive animals.
|
). These animals showed a strong reaction against both bovine and avian tuberculin, but the avian reaction was greater than the bovine reaction. All suspicious cases in the first test were negative in the second confirmative test. Thus, 3.85% of the animals were positive by tuberculin skin test.
Of 125 biopsy specimens obtained from the slaughterhouse, only five samples (from five different animals) yielded positive cultures. In vitro development was slow, i.e., it took 8–21 weeks before growth was visible and all cultures yielded exclusively mycobacteria other than M. tuberculosis (Table 2). None of the samples were positive by Ziehl-Neelsen staining.
|
). Ziehl-Neelsen staining of these samples showed mycobacteria in two mesenteric lymph nodes from both animals and in one lung.
|
). Of three Ziehl-Neelsen–positive samples from the necropsy samples, two were identified by PCR as M. bovis, (both from the same animal), and the other as exclusively mycobacteria other than M. tuberculosis (Table 3). |
DISCUSSION |
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
In previous isolated surveys in Ecuador, the STT and the CTT demonstrated variable prevalences of BTB over time and among various provinces ranging from 0.33% in Tungurahua in 1977 to 4.92% in the Pichincha Province in 2002 (Table 4). Results within the province of Pichincha also differed markedly. A prevalence of 2.80% was observed in Cayambe canton (Andino-Asqui O, unpublished data), which is located in the same province as the region in the present study. However, the results of our study were in sharp contrast to the prevalences of 0.43% (Torres L, unpublished data) and 0.47% (Salazar JC, unpublished data) in a survey of 18 herds in the same area. The 3.85% positive reactors found in the present study are consistent with the 4.92% obtained from 3,089 cattle sampled in Mejía canton on 13 large farms (Cano G, unpublished data).
|
Adams estimated that the sensitivity and specificity of the caudal fold single were 72% and 98.8%, respectively, whereas the sensitivity and specificity of the CTT with bovine and avian PPD was between 68.6% and 95% and 88.8% and 99.9%, respectively.16 The rate of false-negative results is influenced by the time since exposure to environmental strains, immunosuppression, or anergic reactions in the early postpartum period. Desensitization can occur because of too short time intervals between tuberculin skin tests, errors in the procedure, e.g., injection of insufficient tuberculin, use of tuberculins of reduced potency, and test and variability among observers.17 Therefore, in the present study as a precaution, an animal was declared positive when its reaction versus bovine tuberculin was at least 4 mm bigger than the reaction to avian PPD, which is a more rigorous criterion in comparison with the standard used by the United States Department of Agriculture.18
False-positive reactions may be caused by sensitizations as a result of exposure to M. avium, M. paratuberculosis, environmental mycobacteria, and skin tuberculosis caused by slow-growing mycobacteria (M. marinum, M. ulcerans, M. kansasii, M. avium-intracellulare, and M. scrofulaceum) and rapidly growing mycobacteria (M. fortuitum and M. chelonae19). Other factors include the presence of Corynebacterium, Fasciola hepatica, and some Nocardia species, which pose problems in several countries.20 In cattle vaccinated against paratuberculosis, the interpretation of single tuberculin tests might be difficult, but differentiation is possible by the use of the CTT.21 Mycobacterium avium has frequently been recovered from cattle, causing only non-progressive lesions in the mesenteric lymph nodes.22 Sensitization is often caused by exposure to infected domestic or wild birds, and occasionally by exposure to pigs infected with the M. avium-intracellulare-scrofulaceum complex, in which animals show a high reaction to avian PPD, as was observed in five animals from medium-sized farms. Therefore, a positive result in the CCT was restricted to those reactions where bovine tuberculin caused a minimum swelling of 5 mm and differed by at least 4 mm from the swelling caused by the avian tuberculin; the same criterion was used in previous studies in Ecuador (Andino-Ashqui O, unpublished data and Cano G, unpublished data). Although STT and CTT were used in most of the previously performed surveys in Ecuador, the prevalence of aspecific reactions was not clearly mentioned and results referred to the animals positive only for bovine tuberculosis.3
In vitro cultures of 125 specimens from suspected cattle resulted in the isolation of mycobacteria in 5 samples (4%), all of which were mycobacteria other than M. tuberculosis. Thus, suspected postmortem lesions are not necessarily caused by M. bovis as reported by Quinn and others,23 but may be caused by another pathogenic mycobacteria. In a survey in Argentina, bacilli in cattle from slaughterhouses were identified as M. bovis, M. gastri, M. flavescens, M. phlei, and M. triviale.24 In Burundi, Rigouts and others isolated M. bovis from 15 of the 82 cattle sampled; 78% of those isolated grew on Stonebrink medium, which showed the preference of M. bovis for this medium.25 Also identified were mycobacteria other then M. tuberculosis (in descending order of frequency: M. terrae, M. nonchromogenicum, M. intracellulare, M. gordonae, M. sp. rapid grower, and M. paratuberculosis).
The isolation of M. bovis from only 4 of the 29 samples from two positive reactors with a high positive response in the tuberculin test demonstrates the difficulty related to the in vitro culture of this species because the distribution of the bacilli in lesions is not homogeneous.10 Another factor that could have contributed to this is the stage of the disease at the time of sampling. Therefore, this result shows the need for a detailed investigation to avoid false-negative cases. Furthermore, it has been shown that freezing-thawing cycles decrease the viability of mycobacteria.26
Although direct identification of acid-fast organisms by microscopy is fast, it does not identify the M. species.27 In the present study, characteristics that identify M. bovis, such as acid-fast staining, rough morphology of the colony, susceptibility to thiophene-2-carboxylic acid hydrazide, and the absence of niacin production,25 showed the usefulness of in vitro identification, especially when the PCR is not feasible. Microscopic examination by Ziehl-Neelsen staining in 125 specimens from the slaughterhouse and in 29 specimens from the two positives reactors showed the bacilli in only three samples. The low sensitivity of this method is well-known14 because large numbers of organisms (> 104/mL) must be present to make detection reliable.
All positive results obtained by conventional methods have been confirmed by PCR. The advantage of the PCR is that these results were obtained after 48 hours, whereas in vitro identification required 6–8 weeks. In addition, the PCR can determine whether the organisms belong to the M. tuberculosis complex, which requires another week by conventional methods. Since less optimal conditions during transport may have adversely affected the culture results, analysis immediately after sample collection is recommended.
The prevalences observed in the present study confirm the importance of bovine tuberculosis in Mejía canton and stress the need for surveillance at the national level. Use of diagnostic tools such as intradermal tests, in vitro cultures, and PCR will be useful in identifying reactors in herds, increasing epidemiologic information, and differentiating causative species. These tools will increase understanding of the zoonotic consequences, including the possible relation with herd size, and help to implement the most efficient control measures. Whether the PCR as a diagnostic tool is the most useful technique for rapid detection of M. bovis will require further evaluation on a larger sample size. In Ecuador, this technique is still too expensive for routine use. Thus, the STT and CTT in the field and use of in vitro cultures in the laboratory to identify the cause of BTB within a herd is recommended.
Received June 23, 2005. Accepted for publication September 28, 2005.
Acknowledgments: We thank the cattle owners, farm workers, and staff members of the slaughterhouse of the Mejia canton for their assistance and willingness to participate in the study.
Financial support: This study was supported by the Damien Foundation, Belgium, and the Directorate General for Development and Coordination–Institute of Tropical Medicine agreement joint research project 96122.
* Address correspondence to Françoise Portaels, Department of Microbiology, Unit Mycobacteriology, Institute of Tropical Medicine, 155 Nationalestraat, B-2000 Antwerp, Belguim. E-mail: portaels{at}itg.be 
Authors’ addresses: Freddy Proaño-Perez, Jorge Ron, Maria-Augusta Chavez, Richar Rodriguez, and Washington Benitez-Ortiz, International Centre for Zoonoses, Central University of Ecuador, Ciudadela Universitaria, Casillero Postal 17-03-100, Quito, Ecuador, E-mails: fproanio-ciz{at}ac.uce.edu.ec, jron-ciz{at}uce.edu.ec, mchavez-ciz{at}ac.uce.edu.ec, rrodriguez-ciz{at}ac.uce.edu.ec, and wbenitez-ciz{at}ac.uce.edu.ec. Leen Rigouts, Krista Fissette, Anita van Aerde, and Françoise Portaels, Department of Microbiology, Unit Mycobacteriology, Institute of Tropical Medicine, 155 Nationalestraat, B-2000 Antwerp, Belgium, E-mails: lrigouts{at}itg.be, kfissette{at}itg.be, avaerde{at}itg.be, and portaels{at}itg.be. Jef Brandt and Pierre Dorny, Department of Animal Health, Institute of Tropical Medicine, 155 Nationalestraat, B-2000 Antwerp, Belgium, E-mails: jbrandt{at}itg.be and pdorny{at}itg.be.
|
REFERENCES |
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
This article has been cited by other articles:
|
|
 |
|
  L. Durnez, M. Eddyani, G. F. Mgode, A. Katakweba, C. R. Katholi, R. R. Machang'u, R. R. Kazwala, F. Portaels, and H. Leirs First Detection of Mycobacteria in African Rodents and Insectivores, Using Stratified Pool Screening Appl. Envir. Microbiol., February 1, 2008; 74(3): 768 - 773. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
