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Infection and Immunity, August 2003, p. 4544-4553, Vol. 71, No. 8
0019-9567/03/$08.00+0     DOI: 10.1128/IAI.71.8.4544-4553.2003
Copyright © 2003, American Society for Microbiology. All Rights Reserved.

Mycobacterium bovis BCG Scar Status and HLA Class II Alleles Influence Purified Protein Derivative-Specific T-Cell Receptor Vß Expression in Pulmonary Tuberculosis Patients from Southern India

S. Shanmugalakshmi,^1, V. Dheenadhayalan,^1, P. Muthuveeralakshmi,² G. Arivarignan,³ and R.M. Pitchappan^1*

Department of Immunology, School of Biological Sciences,¹ Department of Statistics, School of Mathematics, Madurai Kamaraj University, Madurai 625021,² Government Hospital, Singampunari 630502, India³

Received 12 September 2002/ Returned for modification 19 February 2003/ Accepted 29 April 2003

Top Abstract Introduction Materials and Methods Results Discussion References

 
Purified protein derivative (PPD) RT23-recalled T-cell receptor (TCR) Vß expression was studied in the peripheral blood of 42 pulmonary tuberculosis patients and 44 healthy controls from southern India, a region where tuberculosis is endemic. Forty-eight-hour whole-blood cultures in the presence or absence of PPD-RT23 were set up, and at the end of the culture period total RNA was extracted and cDNA was synthesized. Expression of various TCR Vß families was assessed by using family-specific primers. PPD-specific expression (usage) of TCR Vß families 4, 6, 8 to 12, and 14 was found in more controls than patients. Among the responders (individuals who showed PPD-specific expression), endemic controls had significantly higher responses than the patients had for TCR Vß families 2, 3, 7, 13, and 17. The majority of the patients did not show usage of most of the TCR Vß families, and this was attributed to T-cell downregulation. A four-way nested classification analysis revealed that TCR Vß family 1, 5, 9, 12, and 13 usage in the context of HLA class II high-risk alleles (DRB1*1501, DRB1*08, and DQB1*0601) and Mycobacterium bovis BCG scar status were the determining factors in susceptibility and resistance to tuberculosis. The healthier status of controls was attributed to the wider usage of many TCR Vß families readily recalled by PPD, while the disease status of the patients was attributed to TCR Vß downregulation and the resultant T-cell (memory cell?) unresponsiveness. Host genetics (HLA status) and BCG vaccination (scar status) seem to play important roles in skewing the immune response in adult susceptibility to pulmonary tuberculosis through TCR Vß usage.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Pulmonary tuberculosis, a major infectious disease in the world, was declared a global emergency in 1993 (36). The emergence of multi-drug-resistant Mycobacterium tuberculosis, overcrowded, poor living conditions, and human immunodeficiency virus coinfection have made this a serious health problem in India and other developing countries. The genetic hypothesis that not all individuals infected with the tuberculosis organism develop the disease requires further research (17, 21a). Tuberculosis is a paradigm of conflicts of several factors, including genetics, immunopathology, epidemiology, and host and pathogen diversity (11). Investigating the relative contributions of these factors should lead to better intervention strategies. A major constraint for such analyses is the lack of availability of necessary readouts to evaluate these parameters in the same cohort.

Studies in southern India have identified the high-risk alleles HLA DR2 (DRB1*1501) and DQB1*0601 as predisposing factors for sputum-positive, far-advanced pulmonary tuberculosis (4, 24, 27). Furthermore, expression of the Th2 cytokines interleukin-10 (IL-10) and IL-4 is associated with Mycobacterium bovis BCG scar-negative non-DR2 status in patients (7). It has also been demonstrated that there is a spectrum of immune reactivity (delayed-type hypersensitivity versus serum antibodies) in hospital contacts and healthy individuals in southern India, and this finding has been confirmed in other countries where tuberculosis is highly endemic, such as Indonesia and Brazil (3, 12, 22). However, we are unaware of the epitope specificities of the responses.

Immunologists work under the premise that the epitopes responsible for disease may be different from those affording protection. Thus, the M. tuberculosis epitopes recognized by patients might be different from those recognized by healthy adult controls (immune individuals living in an endemic region). If this is true, the patients must differ from endemic controls in terms of T-cell receptor (TCR) usage for mycobacterial antigens. We should then be in a position to recall this memory in vitro by using a wider antigen, such as the purified protein derivative (PPD) routinely used in skin tests to evaluate sensitization or infection. By restricting the antigen we may not recall global memory (with all the T cells recognizing various epitopes of M. tuberculosis) and may not identify the difference between patients and controls.

An important step in an adaptive immune response is recognition of the peptide major histocompatibility complex by the TCR (8, 25, 37). The TCR repertoire of an individual is the outcome of thymic education in the context of the host major histocompatibility complex. Certain TCR Vß families are more common in individuals with certain HLA class I and class II molecules (repertoire) (28), and HLA identical siblings are more similar in terms of their TCR repertoires than HLA nonidentical siblings (1). The repertoire is also influenced postnatally by environmental exposure to specific and cross-reacting antigens (23). The TCR repertoire of an individual is the memory of experiences of the immune system, including exposure to infectious and innocuous antigens. Thus, it should be possible to identify the immunological memory for a particular antigen by studying TCR Vß expression in the presence and absence of the antigen in vitro.

In this paper we describe the differences between endemic controls and patients (disease status) in terms of PPD-recalled TCR Vß expression. We found a correlation among HLA class II high-risk alleles, BCG scar status, and PPD-specific TCR Vß usage.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Study population. (i) Pulmonary tuberculosis patients. A total of 42 adult pulmonary tuberculosis patients, who were born and brought up in Madurai District in southern India, were enrolled from a state-run public government hospital in Singampunari, Madurai. Ethical clearance from the institutional ethical committee was obtained for the study, and sampling was performed with informed consent. The male/female ratio of the patients was 24:18, and the mean age was 34.7 ± 2.2 years. All the patients showed symptoms of pulmonary tuberculosis with radiological lesions and a compatible clinical picture. The patients were treated with ethambutol, isoniazid, rifampin, and pyrazinamide by following the directly observed therapy protocol implemented in India. The Blood samples were collected during weekly visits to the hospital, and at the time of sampling patients had been treated for various lengths of time (see Table 3).

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TABLE 3. HLA class II alleles, BCG scar status, sputum status, treatment duration, PPD-specific expression of various TCR Vß families, and cytokine expression data in various groups of patients

(ii) Healthy controls. Forty-four healthy endemic controls, all students and staff of Madurai Kamaraj University living in Madurai District, were enrolled for the study. None of these controls had any previous history or symptoms of pulmonary tuberculosis. Their mean age was 26.6 ± 0.76 years, and the male/female ratio was 26:18.

HLA class II typing. HLA DRB1* and DQB1* alleles of the patients and controls were studied by the PCR-sequence-specific oligoprobe method by using primers and probes from the 11th International Histocompatibility Workshop and Conference and the 12th International Histocompatibility Workshop and Conference (2, 18).

Whole-blood cultures. Samples (3 to 5 ml) of peripheral blood were obtained from the patients and controls in heparin vacutainers (455 051; Greiner, Frickenhausen, Germany) and were transferred to the laboratory. Whole-blood cultures were set up on the same day (7). Briefly, 50 µl of whole blood, diluted to a volume of 200 µl with RPMI 1640 medium (31870-025; Life Technologies, Gibco-BRL, Gaithersburg, Md.) and supplemented with 2 mM glutamine (G-1517; Sigma, St. Louis, Mo.), was cultured in quadruplicate for 48 h with or without 20 U of PPD-RT23 in a CO₂ incubator at 37°C in the presence of 5% CO₂ with 95% humidity. After 48 h, the cultures were harvested, and two of the quadruplicate samples were pooled to obtain two aliquots, washed by centrifugation, and frozen at -70°C in lysis buffer.

RNA extraction and cDNA synthesis. Total RNA was extracted from one aliquot of the duplicate aliquots of each sample. RNA was extracted by a single-step acid-phenol-chloroform extraction method (6), and all of the glassware and all of the plastic ware were treated with 0.1% diethyl pyrocarbonate (BDH Laboratory Supplies, Poole, United Kingdom). The RNA was primed with 1 µg of oligo(dT) primer (12- to 18-mer; 27-7858-02; Amersham Pharmacia Biotech, Little Chalfont, Buckinghamshire, United Kingdom) at 65°C for 10 min and cooled immediately on ice. cDNA was synthesized by using Moloney murine leukemia virus reverse transcriptase (E70456Y; Amersham Pharmacia Biotech) in a block heater at 37°C (9), diluted to a volume of 100 µl with diethyl pyrocarbonate water, and stored frozen.

PCR for TCR Vß genes. TCR Vß family-specific primers were synthesized by using the primer sequences described by Hawes et al. (16) and Struyk et al. (32) (Table 1). A total of 24 TCR Vß family-specific 5' primers, one TCR constant-region (TCRC) 5' primer, and one constant-region 3' primer were synthesized by Genosys, Pamisford, Cambridgeshire, United Kingdom. The 3' and 5' constant-region primers were used to amplify the TCRC. The 3' TCRC primer was used with one of the TCR Vß family-specific 5' primers to amplify and identify expression of a given TCR Vß family. Five-microliter portions of the primer pairs were predotted in 96-well plates, stored frozen at -70°C, and used within 1 week of dotting. The PCR mixture (total volume, 20 µl) was dispensed into each well. Each reaction mixture contained 2 µl of cDNA template, 0.5 U of Taq DNA polymerase, each primer at a concentration of 0.2 µM, each deoxynucleoside triphosphate at a concentration of 0.4 mM, 4 mM MgCl₂, and 1x PCR buffer. Twenty-five PCRs were performed for each sample with a Hybaid thermal cycler (Omnigene HBTR3CM; Hybaid Ltd., Middlesex, United Kingdom). The temperature profile was as follows: initial denaturation at 95°C for 1 min; 35 cycles of denaturation at 95°C for 1 min, annealing at 54.5°C for 1 min, and extension at 72°C for 1.5 min; and then a final extension at 72°C for 10 min. The amplified products were electrophoresed at 100 V for 30 min in a 1.5% agarose gel with ethidium bromide. The gels were observed under UV illumination and documented by using a Kodak Digital Science gel documentation and analysis system (Kodak ds EDAS 120; Eastman Kodak Company, Rochester, N.Y.). The TCR Vß products were around 700 bp long, and the TCRC product was 300 bp long (Fig. 1). The band intensities were measured in pixels by using the Kodak Digital Science one-dimensional image analysis software.

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TABLE 1. Primers used to amplify various TCR Vß families and TCRCa

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FIG. 1. Agarose gels showing expression of various TCR Vß families in PHA- or PPD-stimulated, 48-h whole-blood cultures from a healthy control. M, marker X-HaeIII.

Quality control measures. To avoid inter- and intraexperimental variations, RNA extraction and cDNA synthesis for a batch of 10 samples (in the presence and absence of PPD; 20 cDNAs) were performed simultaneously. The PCR, electrophoresis, and measurement of the TCR Vß and TCRC band intensities (in pixels) were all performed simultaneously. The TCRC amplicon served as an amplification control. The net intensities of the TCR Vß bands in no-antigen control and PPD-induced cultures were normalized to the TCRC band intensity of each sample and expressed as percentages of the TCRC band intensity. The antigen (PPD)-specific TCR Vß expression was obtained by subtracting the percentage of the TCRC band intensity for the no-antigen control from the percentage of the TCRC band intensity for the PPD-stimulated cultures. A sample was considered positive for a TCR Vß family if the antigen-specific TCR Vß expression was more than 5% of the TCRC band intensity.

Pilot experiments with whole-blood cultures in two control samples set up in quadruplicate and stimulated with phytohemagglutinin (PHA), PPD, or nothing revealed that PHA induced expression of all TCR Vß families except families 21, 23, and 24 in both donors. The failure with families 21, 23, and 24 was attributed to PCR failure due to primers under the conditions used, and hence these three TCR Vß families were not tested in further experiments. To test the reproducibility of the assay, whole-blood cultures of 10 samples were set up in quadruplicate in the presence of PPD, two cultures were pooled, and the two corresponding cDNAs were synthesized. Expression of TCRC and TCR Vß families was studied in both cDNAs, and the concordance in TCR Vß expression in the two aliquots of the 10 samples was analyzed. Except for TCR Vß families 5, 11, 14, 18, and 19, all of the TCR Vß families showed more than 80% concordance. The lower concordance in some of the TCR Vß families may have been due to difficult primers under the conditions used. The quantitative expression data for various TCR Vß families obtained with the two cDNA aliquots of the 10 samples were concordant. Repeat assays for 11 TCR Vß families gave correlation coefficients of >0.8 (P = 0.003 to P = 0.0001), and repeat assays for six TCR Vß families gave correlation coefficients between 0.7 and 0.8 (P = 0.02 to P = 0.006). In another set of experiments, PCRs with two different samples for TCRC and Vß were performed twice on two different days. Although some of the data were outside the 5% confidence interval, there was a good correlation between the repeat assays (Fig. 2) (R = 0.602, P = 0.003; R = 0.739, P < 0.0001). The results indicated that the methodological approach employed in this study was sufficient to interpret the results obtained.

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FIG. 2. Concordance of repeat assays of TCR Vß expression in response to PHA in two different individuals (A and B). Whole-blood cultures were set up in quadruplicate with 0.4 µg of PHA and harvested after 48 h by pooling two wells. RNAs were extracted, cDNAs were synthesized from both aliquots and pooled, and PCRs for TCR Vß families were performed by using TCR Vß family-specific primers twice on different days. The net intensities for the repeat experiments (in pixels) are compared. The numbers near the data points indicate TCR Vß families. The regression line, 95% confidence interval, and prediction interval are indicated. C, TCR constant region.

Statistical analyses. A paired t test, a Wilcoxon signed rank test, a Mann-Whitney test, and a chi-square test were performed when necessary (29). Log-linear analyses were performed by using BMPD statistical software (13).

Top Abstract Introduction Materials and Methods Results Discussion References

 
Forty-four endemic controls and 42 pulmonary tuberculosis patients were studied for TCR Vß expression in 48-h cultures that were stimulated with PPD or without PPD (no-antigen control). TCRC was expressed in all samples, and the expression was significantly higher in PPD-stimulated cultures than in the no-antigen controls (P < 0.0001). It is for this reason that the expression data were normalized to the corresponding TCRC data (see Materials and Methods). The TCR Vß families expressed in the no-antigen control cultures represented the repertoire, and the difference between the expression in the presence of the antigen and the expression in the absence of the antigen represented the usage.

Comparison of TCR Vß expression in controls and patients. The numbers of controls and patients expressing various TCR Vß families in no-antigen controls were compared (Fig. 3A). We found that 5 to 50% of the samples expressed at least one of the TCR Vß families and that the number of individuals expressing a particular TCR Vß family did not differ significantly between controls and patients except for TCR Vß family 17 (P = 0.048) (Fig. 3A). In the presence of PPD as many as 65% of the controls expressed selected TCR Vß families. Furthermore, more controls than patients expressed many TCR Vß families; eight TCR Vß families (TCR Vß families 4, 6, 8 to 12, and 14) were used by more controls than patients (Fig. 3B).

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FIG. 3. TCR Vß responders in healthy controls and pulmonary tuberculosis patients. The data indicate the percentage of responders in each TCR Vß family. (A) No-antigen control; (B) antigen-specific usage. P values were obtained based on a chi-square analysis with the Yates correction.

A quantitative analysis of the PPD-specific TCR Vß expression data in responders (the individuals who used a TCR Vß family) was also performed (data not shown). The TCR Vß expression levels were significantly higher in responding controls than in responding patients for TCR Vß families 2, 3, 7, 13, and 17 (Mann-Whitney test: P = 0.05, P = 0.019, P = 0.027, P = 0.021, and P = 0.037, respectively).

Thus, there could be two different mechanisms of TCR involvement in protection or the disease process. First, PPD-specific expression of TCR Vß families in more controls than patients indicated that there was antigen-specific immune recognition of selected epitopes in protection. Second, the higher level of selected TCR Vß family expression by controls indicated that there was adaptive immunity and immunological memory in the controls. The absence of these phenomena in patients may indicate that there was TCR Vß downregulation due to antigenic load (25) and, presumably, restricted TCR Vß usage in diseased patients.

The observation described above was further supported by a comparison of TCR Vß expression in the absence and in the presence of antigens in controls and patients. Figure 3 shows that more controls (12 to 26 of the 44 controls) expressed TCR Vß families 1, 5, 6, 9, 10, 11, 13, 14, and/or 18 in the presence of antigen (PPD-specific expression) than in the absence of antigen (1 to 12 of the 44 controls) (P = 0.039, P = 0.009, P = 0.021, P = 0.01, P = 0.019, P = 0.027, P = 0.016, P = 0.005, and P = 0.013, respectively). However, in patients only TCR Vß family 2 showed significant usage (12 and 26 of 42 patients; P = 0.004).

Correlation among HLA high-risk allele status, BCG scar status, and PPD-specific TCR Vß expression. Previous studies in our laboratory showed that there is a high-risk association of HLA DRB1*1501, DRB1*08, and DQB1*0601 with pulmonary tuberculosis (24). Furthermore, cytokine IL-4 expression and IL-10 expression were also associated with disease in non-DRB1*02 patients who were not vaccinated with BCG (7). The TCR Vß expression data presented here were therefore analyzed in the context of the BCG scar status and HLA high-risk allele status.

Tables 2 and 3 show the control and patient samples studied, HLA DRB1 and DQB1 allele data, BCG scar status, PPD-specific expression of various TCR Vß families, and cytokine expression data. The control and patient samples were divided into four groups, based on high-risk allele status and BCG scar status. The number of individuals responding (>5% of TCRC expressed) to a particular TCR Vß family in a group was determined and defined as the number of responders. The total number of TCR Vß families used by each individual was also determined. Among the controls, the total number of responders and the total number of TCR Vß families used were more evenly distributed in all four groups (Table 2). Nonetheless, among the patients, the majority of the responders (9 of 11 responders) expressing more than five TCR Vß families were in the BCG scar-negative high-risk allele group (Table 3). Most of the patients in the other three groups did not respond; only 5 of 25 patients expressed more than five TCR Vß families.

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TABLE 2. HLA class II alleles, BCG scar status, PPD-specific expression of various TCR Vß families, and cytokine expression data for various groups of controls

In order to determine whether chemotherapy has any effect on TCR unresponsiveness in patients, we compared the number of TCR Vß families expressed in patients treated for 50 days (n = 19) and the number of TCR Vß families expressed in patients treated for >50 days (n = 14). This analysis revealed that there was recovery from TCR Vß unresponsiveness in the BCG scar-negative HLA high-risk allele carrier patient group following treatment. Of the 15 patients in this group, 7 were in the group that was treated for 50 days and expressed five or fewer TCR Vß families, 3 were in the group that was treated for 50 days and expressed more than five TCR Vß families, none was in the group that was treated for >50 days and expressed five or fewer TCR Vß families, and 5 were in the group that was treated for >50 days and expressed more than five TCR Vß families (P = 0.0256, as determined by the Fischer exact test). The cytokine data did not exhibit any relationship to the TCR Vß families expressed.

Nested classification analysis of HLA status, BCG status, TCR Vß expression, and disease status. The interactions among the four parameters in question (viz., disease status [D] [healthy versus pulmonary tuberculosis], HLA status [H] [high-risk allele carriers versus non-high-risk allele carriers], BCG scar status [B] [positive versus negative], and TCR Vß family usage status [T] [positive versus negative]) were assessed in a nested classification analysis (13). The number of individuals expressing a particular TCR Vß family in a group of controls and patients was defined as the number of responders in Tables 2 and 3 and used for the analysis.

All possible log-linear models were considered and tested for significance. Four models fit the observed data well (Table 4). TCR Vß families 1, 5, 9, 12, and 13 fit model 1 (TH, TD) well (i.e., interaction of the TCR Vß families with HLA status and with disease status). TCR Vß families 4, 6, 8, 10, 11, 14, and 18 fit model 2 (TD, H, B) well, implying that there were interactions between the TCR Vß families and disease status, which were independent of HLA status and BCG scar status. TCR Vß families 7 and 19 fit model 3 (TH, B, D) well (i.e., interaction between the TCR Vß families and HLA status, independent of BCG status and disease status). TCR Vß families 2, 3, 15, 16, 17, 20, and 22 fit model 4 (T, H, BD) well (i.e., TCR Vß families were independent of HLA status, BCG status, and disease status, although there were interactions between BCG status and disease status). There was no correlation between TCR Vß expression and other parameters, including age, sex, caste, and cytokine expression.

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TABLE 4. Best-fit log-linear models selected by nested classification analyses of HLA status, BCG scar status, disease status, and TCR Vß expression

Top Abstract Introduction Materials and Methods Results Discussion References

 
In the present study we identified the BCG scar status- and HLA class II high-risk allele status-dependent PPD-specific expression of various TCR Vß families in patients and controls. The controls used more TCR Vß families, while the patients used only a few TCR Vß families. Furthermore, the majority of the responders among patients belonged to the BCG scar-negative HLA high-risk allele group.

In the present study carried out in an area in southern India where tuberculosis is endemic, we identified expression of many TCR Vß families in both controls and patients even in the absence of antigen (repertoire). Exposure to various mycobacterial antigens in the environment, atypical mycobacteria, and infection per se may have been the cause of this expression. The expression of TCR Vß families 4, 6, 8 to 12, and 14 in the presence of PPD in significantly more controls than patients (Fig. 3) indicated that there was antigen-specific usage of these TCR Vß families presumably involved in health status. Most of the TCR usage (except usage of family 12) fit model 2 (i.e., TD interaction).

The lack of expression of many TCR Vß families and the restricted usage in patients may have been the result of infection per se. The restricted TCR Vß usage in the peripheral blood of the patients in the present study suggests that there is a focused immune response to some selected epitopes of mycobacterial antigens or downregulation of TCR gene expression due to active suppression. Restricted TCR Vß expression has been reported in multiple sclerosis, human immunodeficiency virus infection, rheumatoid arthritis, and Epstein-Barr virus infection (10, 14, 21, 33). AIDS patients that use one to three TCR Vß families during primary infection deteriorate rapidly, while other patients that use more TCR Vß families deteriorate slowly (26). Patients with chronic hepatitis B show underexpression of TCR Vß families 14 and 15 and expansion of TCR Vß family 7 (5). Reactivation of PPD-specific Th1 lymphocytes with PPD resulted in a concentration-dependent hyporesponsiveness due to an increase in apoptosis of ß, CD4⁺, and ß CD8⁺ cells (30). It is possible that the cells become unresponsive after signaling through their TCR (25). Upon contact with antigen, cells may downregulate their TCR and coreceptors. This may be a consequence of activation under chronic stimulation conditions leading to anergy, a cellular state in which a lymphocyte is alive but fails to exhibit certain functional responses (25). This may be the mechanism of failure of immune surveillance and the resultant disease progression in chronic infections.

Limited TCR diversity with an antigen or infection can be attributed to selection and expansion of certain antigen-specific T cells that become unresponsive in patients (15, 20). The present study showed that this refractory status in patients was determined by BCG scar status and HLA high-risk allele status (Table 3). The downregulation observed might have been due to the enormous amount of M. tuberculosis antigens available in the system due to infection. This possibility was supported by the observation that more TCR Vß families were used by BCG scar-negative and HLA high-risk allele-positive patients that were treated for >50 days than by patients that were treated for <50 days (P = 0.0256). The perturbation in the TCR repertoire and distribution thus seems to be a common feature of various chronic infectious diseases (5, 10, 14, 28, 32, 33).

Investigators in the field of immunology are confronted with dilemmas concerning the site and tissue used for sampling and the antigens that are used. Cells and tissues obtained from the focus of infection may be representative of the disease status and may be the result of the localized immune response. On the other hand, samples from the peripheral blood may indicate the immune surveillance status of the individual. In many previous studies on TCR Vß expression in various disease states the workers have examined the TCR Vß repertoire at the site of disease. In tuberculosis patients, Ohmen et al. identified selective expansion of TCR Vß family 8 in the pleural fluid but not in the peripheral blood (19). In tuberculoid leprosy patients, T cells bearing TCR Vß family 6 are overrepresented in lesions but not in the peripheral blood (34, 35). M. bovis hsp65 peptide-specific T-cell clones express more TCR Vß families 5.1 and J 9 (15, 31). Previous studies of cytokine expression in the peripheral blood of tuberculosis patients and endemic controls have shown that there is PPD-specific enhancement or suppression of gamma interferon and IL-10 following 48 h of culture (7). The reason for the observed PPD-recalled expression of many TCR Vß families in the peripheral blood of more endemic controls than patients in the present study can thus be attributed to the constant exposure to typical and atypical mycobacteria in the endemic environment and the resultant memory. Such exposure has been suggested to be responsible for the spectrum of immune reactivity in endemic controls (3, 12, 22). The exposure and cross-reactivity might have resulted in expansion and persistence of a defined T-cell memory pool, performing immune surveillance. A broader antigen, such as PPD, was thus capable of recalling this global memory (all memory T cells directed towards various epitopes of PPD shared with M. tuberculosis), leading to a better understanding of the factors involved in determining the prevailing adaptive immune status, as indicated in the present study, although we are unaware of the epitope specificity (since it was not the purpose of the study). The use recombinant M. tuberculosis antigens and peptides in various subgroups as reported in this paper may lead to identification of exact epitopes involved in pathogenesis and resistance.

The present study demonstrated that it is possible to identify PPD-recalled memory in the peripheral blood at the TCR Vß usage level and account for the disease status (Table 4). Model 1 of the nested classification analysis revealed that usage of PPD-specific TCR Vß families 1, 5, 9, 12, and 13 along with the HLA class II high-risk alleles was involved in the disease process, while model 2 suggested that TCR Vß families 4, 6, 8, 10, 11, 14, and 18 were also involved in the disease process, although an HLA interaction could not be identified in the cohort used. Model 4 revealed that BCG scar status interacted with the disease status in the presence of TCR Vß 2, 3, 15, 16, 17, 20, and 22, as listed in Table 4. Thus, the disease status was essentially linked to specific TCR Vß usage (models 1 and 2, 12 TCR Vß families) and HLA class II high-risk allele status (model 1, five TCR Vß families). Previous studies have shown that HLA-DR2 status, as well as HLA non-DR2, BCG scar-negative status and IL-10 expression, are associated with pulmonary tuberculosis (4, 7, 24). In the present study we identified a role for TCR Vß usage in tuberculosis susceptibility and protection operating in the context of HLA high-risk allele status and within the parameters of BCG vaccination and environmental exposure.

 
Financial support from the Department of Biotechnology, Government of India, New Delhi (grant BT/PRO281/Med/09/057/96), the Commission of European Communities, Brussels, Belgium (fixed contribution contract C/1-CT93-0079), The Wellcome Trust, London, United Kingdom (grant 061237/Z/00/Z/HH/KO), and the Centre for Scientific and Industrial Research, India (grant CSIR-SRF, Roll no. 112066992) is acknowledged.

We gratefully acknowledge the gift of PPD-RT23 from the BCG vaccine laboratory, Guindy, Chennai, India. Permission from Director of Rural and Public Health, Government of Tamil Nadu (H.Dis.104968/TB/1/97), to carry out this study is acknowledged.

 
* Corresponding author. Mailing address: Department of Immunology, School of Biological Sciences, Centre for Excellence in Genomic Sciences, Madurai Kamaraj University, Madurai 625021, India. Phone: 91-452-2458269. Fax: 91-452-2459181, 139. E-mail: pitchappanrm{at}yahoo.co.uk.

Editor: S. H. E. Kaufmann

Present address: Yerkes Regional Primate Research Centre, Emory University, Atlanta, GA 30329.

Present address: Laboratory of Mycobacterial Diseases, Center for Biologics Evaluation and Research, Food and Drug Administration, Bethesda, MD 20892.

Top Abstract Introduction Materials and Methods Results Discussion References

 

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Infection and Immunity, August 2003, p. 4544-4553, Vol. 71, No. 8
0019-9567/03/$08.00+0     DOI: 10.1128/IAI.71.8.4544-4553.2003
Copyright © 2003, American Society for Microbiology. All Rights Reserved.

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