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The Journal of Immunology, 2006, 176: 3010-3018.
Copyright © 2006 by The American Association of Immunologists

Myobacterium tuberculosis Induces Selective Up-Regulation of TLRs in the Mononuclear Leukocytes of Patients with Active Pulmonary Tuberculosis1

Jung-Su Chang, Jim F. Huggett, Keertan Dheda, Louise U. Kim, Alimuddin Zumla and Graham A. W. Rook2

Centre for Infectious Diseases and International Health, Royal Free and University College Medical School, London, United Kingdom


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Human and mouse studies indicate that TLRs are important in mycobacterial infections. We investigated TLR gene expression in fresh unstimulated blood and bronchoalveolar lavage from patients with pulmonary tuberculosis using a well-validated, real-time PCR. A human splice variant of TLR1, designated hsTLR1, was found in all donors tested. hsTLR1 mRNA lacks exon 2, which is a 77-bp region of the 5'-untranslated region, but contains the same coding sequence as TLR1. Compared with the matched controls, whole blood from patients had increased levels of mRNA encoding TLR2 (p = 0.0006), TLR1 (p = 0.004), hsTLR1 (p = 0.0003), TLR6 (p < 0.0001), and TLR4 (p = 0.0002). By contrast, expression of these TLRs was not increased in bronchoalveolar lavage. An increased level of hsTLR1 mRNA was found in both CD3– (p = 0.0078) and CD4+ cells (p = 0.028), resulting in an increased ratio of hsTLR1 mRNA to TLR1 and to TLR6 mRNA. An in vitro study in THP1 cells suggested that this relative increase in hsTLR1 might be attributable to a direct effect of mycobacterial components because it could be mimicked by mycobacterial preparations in the absence of IFN-{gamma} or T cells and by the TLR1/2 agonist Pam3CysK4. Half-life studies using blood from patients with pulmonary tuberculosis and THP1 cells exposed to Myobacterium tuberculosis in vitro showed p38 MAPK-independent stabilization of mRNAs encoding hsTLR1 and TLR1. We conclude that M. tuberculosis exerts direct effects on patterns of TLR expression, partly via changes in mRNA half-life. The significance of these changes in the pathogenesis of disease deserves further investigation.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Immunity to Mycobacterium tuberculosis (Mtb)3 is associated with Th1 activity and involves TNF-{alpha}, IFN-{gamma}, and IL-12. Neutralization of TNF-{alpha} leads to reactivation of disease (1), and genetic defects of the receptors for IFN-{gamma} or IL-12 lead to increased susceptibility to mycobacteria (2). Until recently it was unclear how infected macrophages are able to link the innate immune response to subsequent T cell-mediated immunity. The discovery of the mammalian TLR addresses this question (3). Ligation of TLRs triggers at least two important signals: 1) a pathway involving MyD88, one of the adaptor proteins shared by all the TLRs, that leads to the activation of the transcription factor NF-{kappa}B, which governs the release of proinflammatory cytokines; and 2) a second signal, which can be MyD88-dependent or MyD88-independent, drives the maturation of APCs and increases the expression of MHC molecules, costimulatory molecules, CD40, and the chemokine receptor of CCR7. These two signals are believed to be essential for the initiation of T cell-mediated immunity (4).

Eleven members of the TLR family have been identified, and each activates a unique downstream gene profile (5). TLRs recognize conserved pathogen-associated molecular patterns (3). Two members of the TLR family, TLR2 and TLR4, have been shown to mediate Mtb-induced intracellular signaling by purified ligands or live bacilli (6, 7). Several components of Gram-positive bacteria are recognized by TLR2, whereas the LPS of Gram-negative bacteria is usually recognized by TLR4 (8). Alternative dimerization of TLR2 with TLR1 or TLR6 allows TLR2 to recognize subtle differences in lipid configuration (9, 10). In summary, TLR2/TLR6 heterodimers recognize the lipoteichoic acid of Gram-positive bacteria and the peptidoglycan and soluble tuberculosis factors of Mtb (9, 11, 12, 13). Agonists for TLR1/TLR2 heterodimers include the 19-kDa lipoprotein of Mtb, the synthetic lipopeptide Pam3CysK4, and ara-lipoarabinomannan of mycobacteria (10, 14, 15, 16, 17). Mycobacterial lipoarabinomannan is also a ligand for the CD14/TLR2 receptor complex (18) as well as for the recently identified lectin dendritic cell-specific ICAM-3 grabbing nonintegrin designated DC-SIGN (19, 20). The 24-kDa LprG of Mtb is a ligand for TLR 2, although it is not currently known whether TLR1or TLR6 is also involved (21).

Early work using human TLR2 or TLR4 overexpressed in cell lines suggested that viable Mtb bacilli activate cells via both TLR2 and TLR4 (7). Subsequent studies in mice with inactivated TLR genes showed that TLR2 is important in controlling (22) and surviving (23) Mtb infection. However, some reports suggested that TLR4 is also important for survival (24, 25, 26), whereas other reports argued that the importance of TLR4 may depend on the dose of Mtb used for challenge (27) or the choice of mouse strain (28). Human studies show that polymorphisms of TLR2 or TLR4 can result in increased susceptibility to microbial infections, possibly by influencing Th1/Th2 balance (15, 29, 30, 31, 32). Interestingly, a mutation (R677W) in the intracellular domain of TLR2 has been linked with lepromatous leprosy, which shows a Th2-like profile (31). These patients had diminished cellular responses to both Mtb and Mycobacterium leprae (33).

Given the critical role of TLRs in innate immunity and initiation of the appropriate adaptive response, the regulation of TLR expression is likely to be an important determinant of the clinical outcome of Mtb infection. Moreover, the presence of Th1 or Th2 cytokines can modulate the expression and also the activation level of TLR1 and TLR2 in response to M. leprae (15). Interestingly, Fenhalls et al. (34) reported that the pattern of TLR expression in tuberculous lung granulomas correlated with the presence or absence of immunohistologically detectable IL-4. When examining the TLR distribution in granulomas, it was shown that TLR1, TLR2, and TLR4 were expressed in both immune cells (e.g., lymphocytes and myeloid cells) and nonimmune cells (e.g., epithelium, type II pneumocytes, and pericytes), whereas TLR9 was restricted to immune cells. TLR3 and TLR 5 were exclusively expressed on myeloid cells. This immunohistological study (34) is the only ex vivo study of TLR expression in tuberculosis (TB) of which we are aware, but in view of the increasing evidence for the importance of IL-4 (35), it raised the possibility that changes in TLR expression might represent useful markers of the immunological status of patients and their contacts. Therefore we used a well-validated, real-time PCR to measure the expression of mRNA encoding TLRs in the fresh unstimulated whole blood and bronchoalveolar lavage (BAL) of tuberculosis patients and matched controls. First, we demonstrated striking changes in the expression of TLRs in the peripheral blood of patients but not in the lavage cells, suggesting a possible down-regulation of TLR expression by cytokines at the site of the infection. Second, a novel human splice variant of TLR1 (designated hsTLR1) showed the greatest response to Mtb both in vivo and in vitro, resulting in a changing ratio of hsTLR1 relative to TLR1 and to TLR6. A study of mRNA half-life in whole blood from patients and in THP1 cells exposed to Mtb in vitro showed p38 MAPK-independent stabilization of mRNAs encoding hsTLR1 and TLR1. Posttranscriptional stabilization of hsTLR1 and TLR1 mRNA after exposure to Mtb might account for increased expression in whole blood but may not be sufficient to explain the increase in mRNA encoding hsTLR1 relative to TLR1.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Patients and samples

Ten patients with pulmonary TB were recruited at the Royal Free and Middlesex Hospitals, London (U.K.). All patients yielded positive cultures from sputum or alveolar lavage fluid, were negative for Abs to HIV, and responded clinically to anti-TB treatment. Informed consent was obtained from all patients, and the study was approved by the relevant ethical review committee. The details of the recruitment criteria of subjects can be seen in our previous published study (36). Controls were matched to the cases for age (within 4 years), gender, and ethnicity. To exclude latent Mtb infection in the controls, T cell IFN-{gamma} ELISPOT responses to early secreted antigenic target 6 and culture filtrate protein 10 peptide pools were determined (T-SPOT.TB; Oxford Immunotec).

Whole blood collection, cell separation, and flow cytometry

Whole blood (2.5–20 ml) was taken before or within the first 2 wk of anti-TB treatment, and 2.5 ml was immediately transferred from the patient into PAXgene blood RNA tubes (PreAnalytix; Qiagen) to fix the mRNA profile. The remaining blood was used for additional experiments (cell subpopulation studies, IFN-{gamma} and IL-4 ELISPOT assays, and mononuclear cell culture).

PBMCs were separated from heparinized whole blood by density gradient centrifugation. For cell subpopulation studies, the enrichment of CD3+, CD3+CD4+, and CD3+CD8+ and the depletion of T cells were performed using Ab-based density centrifugation reagents according to the manufacturer’s directions (RosetteSep; StemCell Technologies). Cell purity was confirmed by flow cytometry after staining and fixing cells with anti-CD4-FITC, anti-CD8-PE, and anti-CD3-PerCP Abs (BD TriTEST; BD Biosciences). Mean cell purities of the relative fractions were >95, 99, 93, and 90% for CD3+, CD3–, CD4+, and CD8+ fractions, respectively, and were determined by flow cytometry using the CellQuest 3.3 (FACScan; BD Biosciences).

Bronchoalveolar lavage

BAL was undertaken under local anesthetic and midazolam sedation in patients (n = 15) and controls (n = 5). To obtain BAL fluid, 0.9% saline was instilled into a radiologically involved lung segment using 180 ml of saline in 60-ml aliquots. In control donors the right middle lobe was lavaged.

Cell culture

The human myelomonocytic cell line THP1 (American Type Culture Collection (ATCC)) was cultured in RPMI 1640 medium supplemented with 2 mM L-glutamine, 100 U/ml penicillin, 100 µg/ml streptomycin (all from Invitrogen Life Technologies), and 10% FBS at 37°C with 5% CO2. For the induction of cell differentiation, cells (2 x 105/ml) were initially propagated in 250-ml flasks (Nunc) for 2 days. The cells were then treated with 1.2% DMSO (Sigma-Aldrich) for 24 h to differentiate them into macrophages (37). DMSO was removed by washing it off twice with wash medium (RPMI 1640 with penicillin-streptomycin), and cells were resuspended to 2 x 106/ml in culture medium. Cell viability was assessed by trypan blue exclusion. For cell stimulation, 2 x 106/ml differentiated cells were incubated with different concentrations of mycobacterial Ags (90 µg/ml sonicated Mtb (sMtb), 300 µg/ml sonicated M. vaccae (SMv), and 4.8 µg/ml LPS (Sigma-Aldrich)) and PBS as negative control, and cells were harvested at 6, 18, 24, and 48 h posttreatment. In subsequent experiments, differentiated THP1 cells were treated with 100 µg/ml Pam3CysK4 (Bachem) or 90 µg/ml sMtb for various times. The concentrations of Ags used in the experiments were optimized in pilot experiments using IL-12 p40 cytokine release as the readout.

RNA stability

Heparinized blood was taken from controls and TB patients within the first 2 wk of anti-TB treatment and was cultured with or without 5 µg/ml actinomycin D (ActD; Sigma-Aldrich) in 24-well plates (Nunc) at 37°C in 5% CO2. At time 0 and after 1, 2, and 4 h, blood was transferred into PAXgene blood RNA tubes (PreAnalytix; Qiagen). To measure mRNA half-life in THP1, differentiated cells (2 x 106/ml) were incubated with or without 90 µg/ml sMtb for 6 h. Both stimulated and unstimulated cells were treated with or without 10 µg/ml ActD, and cells were harvested at the indicated times. (Optimal concentrations of ActD for use in blood and THP1 cell cultures had been titrated in pilot experiments.) The effect of ActD on blocking transcription activity was assessed using Myc proto-oncogene (c-myc) transcription factor p64 as positive control (38). To check whether changes in mRNA half-life were p38-MAPK-dependent, differentiated THP1 cells were treated with 90 µg/ml sMtb for 6 h. Cells were then treated with 10 µg/ml ActD with or without 5 or 10 µM p38 antagonist (SB202190) (Calbiochem). Osteogenic differentiation factor (SOX 9) gene expression was used as a positive control for the effects of SB202190, because the half-life of its mRNA is known to be regulated by p38 in THP1 cells (39).

Suspensions of living mycobacteria

Mtb H37Rv (ATCC no. 25618) and the fast growing environmental saprophyte M. vaccae (National Collection of Type Cultures no. 11659) were grown in Middlebrook 7H10 agar (Difco) containing 10% v/v oleic acid/albumin/dextrose/catalase supplement (BD Biosciences) in the category 3 laboratory. Colonies were transferred into sterile screw-top Eppendorf tubes that contained deionized water with 0.1% v/v tyloxapol and four sterile 1.5- to 2-mm glass beads. Mycobacteria were disaggregated by vigorous vortexing with glass beads for 1–2 min, and the tubes were left standing for 5 min to allow large aggregates to settle. The supernatant suspension was then removed, centrifuged at 1200 rpm for 2 min, and diluted (1/10) in formaldehyde for counting. Ten microliters of the suspensions was added to improved Neubauer hemocytometers, and organisms were allowed to settle for at least 30 min at room temperature. A direct microscopic count was performed to determine mycobacterial concentration.

Infection of PBMCs

PBMCs from five healthy control donors were separated from heparinized blood (50 ml) by Ficoll density gradient centrifugation, and cells were reconstituted at 2 x 106 cells/ml. PBMCs were infected with living mycobacteria at a dose of ~1 organism per macrophage (~10% monocytes in PBMC). Cells were cultured in RPMI 1640 supplemented with 5% heat-inactivated human AB serum and 1% L-glutamine without antibiotics at 37°C and 5% CO2 in a category 3 laboratory and harvested at 18, 24, 48, and 66 h posttreatment. The viability of mycobacteria was assessed by culturing the diluted bacteria on Middlebrook 7H10 agar.

RNA isolation, quality assessment, and reverse transcription

Total RNA was extracted using PAXgene blood RNA tubes (Qiagen) or RNeasy mini kit (Qiagen). All samples were treated with DNase (Qiagen). Cells were harvested at the indicated time points. After centrifugation, pellets were lysed immediately to prevent unwanted changes in RNA profile. The RNA template was qualitatively assessed and quantified using an Agilent 2100 Bioanalyzer (RNA 6000 Nano LabChip kit; Agilent). Contamination of samples with genomic DNA or degraded RNA was assessed on electrophoretic traces or gel bands. A fixed amount of total RNA (1 µg/reaction for culture experiments, 25 ng/reaction for whole blood and cell subpopulation studies, and 500 ng/reaction from BAL) was incubated with 1.5 µM oligo(dT) primer (Promega) at 75°C for 5 min to denature RNA. The reverse transcription master mix for the culture experiments and BAL was (10x reverse transcriptase buffer, 0.5 mM dNTP, 10 U of RNase inhibitor, 4 U of Omniscript reverse transcriptase, and RNase-free water) (Qiagen). Sensiscript reverse transcriptase was used for the whole blood and cell subpopulation study (10x reverse transcriptase buffer, 0.5 mM dNTP, 10 U of RNase inhibitor, 10 U of Sensiscript reverse transcriptase, and RNase-free water) (Qiagen). Denatured RNA was added to 20 µl of reverse transcriptase reaction mix and incubated at 37°C for 1 h followed by 93°C for 5 min to denature the enzyme. The cDNA was aliquoted and used for quantitative PCR.

Quantitative RT-PCR

Primers and probes for each gene studied were designed using the primer 3 web site (http://frodo.wi.mit.edu/cgi-bin/primer3/primer3_www.cgi), and the details of primer concentration and sequences are shown in Table I. Primers were synthesized by MWG Biotech, and probes were purchased from Sigma-Aldrich. All of the designed amplicons were <150 bp to ensure high amplification efficiency (40). The primer (probe) concentration and the annealing temperature of each gene were optimized. The human acidic ribosomal protein HuPO was selected as reference gene in this study due to its stability in response to Mtb treatment (41, 42). RNA expression profiles of both target and reference genes were performed using the RotorGene (Corbett Research) as 12.5 µl/PCR (optimizing primers per probe concentration, 1 µl of cDNA; QuantiTect RT-PCR Master Mix buffer (Qiagen)). Reporters were either the nonspecific DNA binding dye SYBR Green using the QuantiTect SYBR Green Master mix (Qiagen) or a specific hydrolysis probe with QuantiTect Probes Master Mix (Qiagen). Standards for gene quantification were 10-fold dilutions of plasmid DNA. The product sizes of plasmid DNA were confirmed by electrophoresis in 1.8% Tris-acetate-EDTA agarose gel (SYBR Green) and a 1-kb ladder (Promega). Inserts of TLR1 and hsTLR1 were further confirmed by sequence analysis.


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  		Table I. Primer and probe sequences used to quantify gene expression by real-time PCR

 
Data analysis

The Mann-Whitney U test was used for the analysis of differences in the clinical study. Time-course experiments were analyzed by two-way ANOVA with Bonferroni posttests and correction. One-phase exponential decay was used to analyze the RNA degradation rate. Analysis was conducted using GraphPad Prism 4.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
Discovery of a novel splice variant of TLR1, designated hsTLR1

During the initial screening, a human splice variant of TLR1, designated hsTLR1, was discovered in THP1 cells using the TLR1-1-3 primer (Table I). Agarose gel electrophoresis suggested that the larger product was the intended amplicon (142 bp), whereas the second amplicon was <100 bp (Fig. 1A). Sequence information confirmed that the smaller amplicon differed by the excision of exon 2, which forms part of the 5'-untranslated region (UTR) (Fig. 1B). Specific primers were designed to distinguish TLR1 from the novel splice variant of TLR1 (Table I). Subsequent studies of cell distribution indicated that hsTLR1 was expressed in PBMCs from all donors tested (n = 10) as well as in CD19+ cells (n = 5, data not shown), although the baseline expression level of hsTLR1was lower than that of TLR1 in controls. We then investigated the effect of excision of exon 2 on the secondary structure of the RNA as determined by Mfold using a minimum free energy prediction (http://www.bioinfo.rpi.edu/~zukerm/rna/). Exon 2 can fold upon itself (data not shown). However, no key structures were identified using the RNA Families Database of Alignments and Covariance Models (http://www.sanger.ac.uk/Software/Rfam/index.shtml). We cloned and sequenced the coding region of hsTLR1 and confirmed that it is the same as that of TLR1 (data not shown).


Figure 1
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  		FIGURE 1. Discovery of a splice variant of TLR1 in THP1 cells. Forward and reverse primers to TLR1 cDNA bind within exon 1 and 3, respectively, to generate a 142-bp amplicon. A, Agarose gel electrophoresis of RT-PCR products revealed two products: the larger molecule is the target amplicon (142 bp), and the smaller molecule is < 100 bp. B, Sequence alignment of the two amplicons demonstrated that the smaller molecule lacks exon 2.

 
Effects of tuberculosis infection on levels of expression of TLR mRNA in whole blood

Compared with the matched controls, whole blood from patients had significantly increased levels of mRNA encoding TLR2 (p = 0.0006), TLR1 (p = 0.004), hsTLR1 (p = 0.0003), TLR6 (p < 0.0001), and TLR4 (p = 0.0002) (Fig. 2). By contrast, TLR7 and TLR9 mRNAs were not increased in cells from patients (TLR3, TLR5, and TLR8 were not studied). Up-regulation of mRNA encoding TLR2 and TLR2 heterodimerization partners (hsTLR1, TLR1, and TLR6) (Fig. 2) resulted in altered ratios of TLR2 mRNA to mRNAs for TLR1, hsTLR1, and TLR 6 in whole blood, although these changes were not significant (Table II).


Figure 2
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  		FIGURE 2. Expression of mRNA encoding TLR genes in fresh unstimulated whole blood from patients with progressive TB compared with healthy matched controls (n = 10). Whole blood (2.5 ml) was taken before or within the first 2 wk of anti-TB treatment and transferred immediately into PAXgene blood RNA tubes to fix the mRNA profile. Data were normalized relative to 106 copies of the human acidic ribosomal protein HuPO (huPO). Expressions of TLR2 (A) and TLR4 (B) were strikingly up-regulated in patients (p = 0.0006 and p = 0.0002, respectively, compared with control donors). TLRs that form heterodimers with TLR2 were also up-regulated in the whole blood of patients. C, TLR1 (p = 0.004). D, hsTLR1, the splice variant of TLR1 (p = 0.003). E, TLR6 (p < 0.0001).

 

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  		Table II. Ratios of TLR2 mRNA to mRNAs for TLR1, hsTLR1, and TLR 6 in whole blooda

 
Expression of TLR genes in cell subpopulations

Fresh PBMCs were separated into CD3–, CD3+, CD3+CD4+, and CD3+CD8+. Purity of the fractions was confirmed by flow cytometry. Specific surface Ab staining indicated 99, 95, 93, and 90% purity, respectively. There was an increased level of hsTLR1 mRNA in CD3– cells (mostly non-T cells) from patients (p = 0.0078), as well as a modest increase in the CD4+ population (p = 0.028) (Fig. 3). Expression of TLR7 showed a small increase in the CD3+ population (p < 0.05; data not shown). Up-regulation of the mRNA encoding of hsTLR1 in CD3– cells of patients (Fig. 3) resulted in an increased ratio of hsTLR1 to TLR1 (p = 0.017) and to TLR6 (p = 0.013) (Table III). Similarly, although the increase in expression of hsTLR1 in the CD4+ population of patients was small (p = 0.028), it resulted in a marked increase in the ratio of hsTLR1 to TLR1 (p = 0.004) and to TLR6 (p = 0.0031) in this population (Table III).


Figure 3
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  		FIGURE 3. Expression of mRNA encoding TLR in cell subpopulations from fresh unstimulated whole blood from patients and controls. Fresh PBMCs were separated into CD3–, CD3+, CD3+CD4+ (CD4+), and CD3+CD8+ CD8+), and the purity of fractions was confirmed by flow cytometry (99, 95, 93, and 90% purity, respectively). There was increased mRNA for hsTLR1 in the CD3– population from patients (p = 0.008) (A). The mRNA for hsTLR1 was marginally increased in the CD4+ population from patients (p = 0.028) (B).

 

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  		Table III. Ratios of hsTLR1 mRNA to mRNAs for TLR1 and TLR6 in cell subpopulationsa

 
Expression of mRNA encoding TLR in BAL cells

In contrast to the large increases in expression of TLR in whole blood from tuberculosis patients, the levels of expression in BAL were not different from those seen in lavage from matched control donors (Fig. 4).


Figure 4
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  		FIGURE 4. Expression of mRNA encoding TLR in BAL. Fresh BAL taken from patients (n = 15) and controls (n = 5) were immediately placed in mRNA-stabilizing buffer. mRNA levels were assayed exactly as described for whole blood. In contrast to the results with whole blood (Fig. 2), there was no increase relative to BAL from control donors. Results are shown for TLR1 (A), hsTLR1 (B), TLR2 (C), TLR4 (D), TLR6 (E), and TLR7 (F).

 
Effects of Mtb and Pam3CysK4 on expression of hsTLR1 in vitro

We then tested the hypothesis that the increases in TLR expression and the relative increase in hsTLR1 might be attributable to a direct effect of mycobacterial components rather than to an indirect effect requiring IFN-{gamma} or other T cell products. Treating THP1 cells with sMtb (90 µg/ml) or sMv (300 µg/ml) resulted in significantly increased mRNAs encoding hsTLR1 and TLR1 and a significantly increased ratio of hsTLR1 to TLR1 (data not shown).

Because mycobacteria contain components that trigger TLR2/TLR1 heterodimers such as lipoarabinomannan and the 19-kDa lipoprotein of Mtb, we next tested whether a known defined trigger of TLR2, Pam3CysK4, would also increase expression of these TLRs and increase the ratio of hsTLR1 to TLR1. Incubation with Pam3CysK4 led to increased expression of both hsTLR1 and TLR1 at 18 and 24 h, (Fig. 5, A and B) but only TLR1 was increased at 48 h. In the presence of sMtb, the increased expression of hsTLR1 persisted at 48 h. All of the stimuli, including Pam3CysK4, led to an increased ratio of hsTLR1 to TLR1 at 18 (p < 0.01) and 24 h (p < 0.01 with sMtb; p < 0.001 with Pam3CysK4), but only sMtb caused a persistent increase in this ratio at 48 h (p < 0.001) (data not shown).


Figure 5
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  		FIGURE 5. Effects of sMtb and Pam3CysK4 on the expression of mRNA encoding hsTLR1 and TLR1 in THP1 cells. Cells cultured with sMtb had increased hsTLR1 gene expression at 18, 24, and 48 h (A), whereas Pam3CysK4 induced hsTLR1 expression at 18 and 24 h but not at 48 h. By contrast, both sMtb and Pam3CysK4 caused increased expression of TLR1 at 18, 24, and 48 h (B). Results are from one of the repeat experiments and are presented as means ± SD. (**, p < 0.01; ***, p < 0.001)

 
Stability of hsTLR1 mRNA and the p38 pathway

The increase in hsTLR1 relative to TLR1 driven by Mtb components suggested that hsTLR1 was more sensitive to Mtb than TLR1. This phenomenon could be explained either by immune cells altering their splicing ratio or by specific changes in the respective RNA half-lives when in contact with Mtb. The half-lives of mRNAs encoding hsTLR1 and TLR1 were examined in ex vivo blood from TB patients (n = 7) and control donors (n = 4). Fig. 6A shows that the half-lives of TLR1 and hsTLR1 mRNA were similar in whole blood from the control donors (hsTLR1 half-life of 68.07 min vs TLR1 half-life of 61.49 min), but both half-lives were prolonged in blood from TB patients. Because sMtb and Pam3CysK4 increase expression of hsTLR1 mRNA in vitro, we asked whether exposure to sMtb would increase the half-life of hsTLR1 mRNA in THP1 cells. The steady-state mRNA half-lives of hsTLR1 and TLR1 were similar in THP1 cells, namely 154.3 min vs 180.6 min, respectively. Treatment with sMtb delayed RNA degradation of mRNA encoding hsTLR1 and TLR1 at 240 min (p < 0.001) and 360 min (p < 0.001) compared with unstimulated wells (Fig. 6B). sMtb-induced stabilization in RNA encoding hsTLR1 also led to increase relative stability of hsTLR1 over TLR1 at 240 min (p < 0.01).


Figure 6
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  		FIGURE 6. RNA stability in whole blood and THP1 cells. Whole blood cells from control donors (n = 4) and TB patients (n = 7) were incubated with 5 µg/ml ActD, and cells were harvested at the indicated time points. There was a trend toward prolonged stability of hsTLR1 and TLR1 mRNA in samples from patients (A). THP1 cells were cultured with or without sMtb (90 µg/ml) for 6 h. ActD (10 µg/ml) was then added to each well. Incubation with sMtb delayed degradation of mRNAs encoding TLR1 and hsTLR1 at 240 min (p < 0.001) and 360 min (p < 0.001) compared with unstimulated wells (B). The effect of sMtb on the stability of hsTLR1 mRNA was greater than its effect on TLR1 mRNA at 240 min (p < 0.01). Results are from one of the repeat experiments and are presented as means ± SD. Two-way ANOVA with Bonferroni’s correction was performed.

 
MAPK p38 is involved in the stabilization of mRNAs encoding a number of inflammatory cytokines such as TNF-{alpha} and IFN-{gamma} (43, 44). Therefore, we examined the role of the p38 MAPK pathway in posttranscriptional regulation of TLR1 and hsTLR1 mRNA. The addition of the p38 MAPK inhibitor SB202190 caused accelerated degradation of mRNA encoding SOX-9, the half-life of which is known to be regulated in this way, but had no effect on the stability of mRNA encoding TLR1 or hsTLR1 (data not shown).

The effects on TLR mRNA levels of infecting PBMC with living mycobacteria in vitro

We investigated whether the effects seen in the ex vivo studies with patients’ cells could be reproduced by infecting PBMC from normal donors with living mycobacteria at a dose of ~1 organism per macrophage. Expression of TLR2 increased in PBMC after 48 h in culture even in the absence of mycobacteria. However, there was a significant further increase in TLR2 mRNA in the presence of H37Rv at 48 h (p < 0.01) and 66 h (p < 0.01) (Fig. 7A). H37Rv did not cause significant changes in the expression of TLR4 (Fig. 7B) or TLR7 (data not shown). Expression levels of the coreceptors of TLR2, TLR1, and TLR6, were also not changed (Fig. 7, C and E), but there was an increase in expression of hsTLR1 after 66 h of culture with H37Rv (Fig. 7D, p < 0.05). The environmental saprophyte, M. vaccae, caused a distinct pattern of changes with inhibition of TLR4 at 48 h (p < 0.001) and 66 h (p < 0.01) (Fig. 7B). Suppression of TLR1 (Fig. 7C) and an early increase in expression of hsTLR1 (Fig. 7D) were also observed in wells incubated with M. vaccae, but these changes were not significant.


Figure 7
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  		FIGURE 7. Kinetics of expression of TLR in human PBMCs in response to incubation with living Mtb H37Rv or M. vaccae. Incubation with H37Rv caused increased expression of TLR2 at 48 h (p < 0.01) and 66 h (p < 0.01) (A) and a borderline increase in hsTLR1 at 66 h (p < 0.05) (D) compared with control wells. By contrast, H37Rv did not cause any change in expression of TLR4 (B) or expression of potential TLR2 heterodimerization partners TLR1 and TLR6 (C and E). M. vaccae induced a distinct pattern of changes, with suppression of TLR4 at 48 h (p < 0.001) and 66 h (p < 0.01). Results are from healthy control donors (n = 5) and are presented as means ± SD.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
Ex vivo studies that involve no in vitro stimuli and minimal laboratory manipulation provide a "snapshot" of the situation in vivo and have the potential to suggest surrogate markers for clinical application. We have previously used this approach successfully to study the relevance of IL-4 and the inhibitory splice variant of IL-4, IL-4{delta}2, in TB patients and their contacts. The data presented here show that expression of mRNAs encoding TLR1, hsTLR1 (a novel splice variant of TLR1), TLR2, TLR4, and TLR6 are all strikingly increased in fresh unstimulated whole blood from TB patients as compared with their levels in cells from matched controls. We cannot draw conclusions about disease specificity, because no other disease group was studied. The findings might, as discussed below, be expected in other infections associated with triggers of TLR2. By contrast, there was no such increase in the expression of mRNA encoding TLR in BAL from patients. This finding might be explained by our previous observation that levels of IL-4 mRNA are increased more in the BAL of TB patients than in their blood. The copy number of IL-4 mRNA was almost 100 times higher in the BAL from the patients studied here as compared with BAL from matched controls (36). These observations are in agreement with the only other ex vivo study of TLR expression in TB of which we are aware. Fenhalls et al. (34) used immunohistochemical analysis of lung granulomas to show detection of TLR1, TLR2, TLR3, TLR4, TLR5, and TLR9, but levels of TLR2 and TLR4 were lower in those granulomas in which IL-4 mRNA was detected. IL-4-mediated down-regulation of TLR2 expression in monocytes and dendritic cells in vitro was reported by Krutzik et al. (15). It will clearly be of some interest to examine macrophages in BAL from TB patients for the patterns of activation attributable to Th1 cytokines or Th2 cytokines (so-called alternative macrophage activation) (45).

In freshly drawn blood cells from tuberculosis patients, the TLR mRNA showing the greatest relative increase was found to encode a splice variant of TLR1 that we have designated hsTLR1. We show here that hsTLR1 mRNA lacks exon 2, which is a 77-bp region of the 5'-untranslated region. We cloned and sequenced the coding region of hsTLR1 and confirmed that it is the same as TLR1 (data not shown). We then investigated the secondary structure of exon 2 using the Mfold web server and demonstrated that this sequence could form significant secondary structure. hsTLR1 mRNA was expressed in myeloid cells, T cells (CD4+ and CD8+), and CD19+ B cells of all donors tested (n = 10, except for CD19+, where n = 5) and was induced in a human monocytoid cell line (THP1 cells) by the TLR1/2 ligand Pam3CysK4. A half-life study in whole blood and in THP1 cells exposed to Mtb in vitro showed stabilization of mRNAs encoding hsTLR1 and TLR1, which was independent p38 MAPK. The half-lives of the two variants increased in parallel, so changes in mRNA half life might not explain the increase in hsTLR1 relative to TLR1 and to TLR6 seen in T cells and non-T cells from tuberculosis patients.

Steady-state mRNA levels represent a balance between message stability and the rate of gene transcription. Messenger RNAs for many molecules involved in inflammation and stress responses are normally unstable, presumably to allow rapid regulation of responses to environmental changes (46). Fan et al. (47) reported that the expression of TLR4 is destabilized after challenge with LPS in a murine model, and both transcription activity and RNA stability contributed to the expression level of TLR4. However the authors did not define the underlying mechanisms that account for the posttranscriptional regulation of the murine TLR4 gene. Several components are known to influence RNA stability: the 5' cap, the 3' poly(A) tail, adenosine- and uridine-rich elements within the 3'-UTR, sequences within the 5'-UTR, and sometimes motifs within the protein-coding sequence itself (48). Tandem repeats of AU-rich elements, AUUUA, in the 3'-UTR regions are associated with a short mRNA half-life, but the instability is countered by phosphorylation of RNA-binding molecules via the p38 pathway when the cell becomes activated (49, 50). However, our data do not support a role for p38 MAPK-dependent stabilization of TLR1 mRNA in Mtb-exposed cells, so the underlying mechanism remains unknown. Moreover, because the stability of the mRNA encoding TLR1 and hsTLR1 increases similarly in TB and in cells exposed to Mtb in vitro, changes in half-life are unlikely to explain the increase in hsTLR1 relative to TLR1 seen in this work.

The size of the increases in expression of TLR in peripheral blood, remote from the clinically involved lung tissue, is surprising. This finding might indicate that expression of TLR1, hsTLR1, TLR2, TLR4, or TLR6 at the mRNA level is sensitive to increased circulating levels of proinflammatory cytokines or to circulating mycobacterial components. The latter possibility was supported by the observation that the infection of PBMC from normal ELISPOT negative donors with live mycobacteria could mimic some of the effects seen in the ex vivo studies. The finding that sMtb or sMv could cause the similar effects in THP1 cells in the absence of T cells led us to test a defined TLR2 agonist, Pam3CysK4. This molecule potently up-regulated TLR2 and TLR1 and also increased the ratio of hsTLR1 to TLR1, as found in the cell subsets in the ex vivo studies. Interestingly, Pam3CysK4 replicated the effects of M. vaccae but failed to cause the sustained increase in ratio of hsTLR1/TLR1 that was seen at 48 h in the presence of sMtb.

The strikingly increased ratio of hsTLR1 to TLR1 and TLR6 induced by the mycobacteria and by Pam3CysK is interesting. TLR1 and hsTLR1 share the same protein coding sequences. Nevertheless, splice variants of noncoding regions of other genes have been implicated in the regulation of translational efficiency (51, 52, 53). Therefore, a switch to hsTLR1 could modulate relative levels of TLR1 and TLR6 and thus alter the composition and relative frequency of the TLR2/TLR1 or TLR2/TLR6 heterodimers. This issue was not addressed here. However there is evidence that TLR2 function is important for immunity to mycobacteria in mice and humans. Defective TLR2 activation during the infection can favor Th2 cytokines (54, 55) and could be involved in the existence of both Th1 and Th2 cytokines at the site of infection in TB patients (34, 36). Therefore, we hypothesize that the outcome of TLR2 activation may depend on the ratio of hsTLR1 to TLR1 and to TLR6. Further work will be needed to explore this hypothesis.

In conclusion, the present study indicates that the expression of several TLR genes is strikingly increased in ex vivo blood samples from patients with progressive TB. Such increased expression was not seen in BAL samples from the same donors. This finding might indicate down-regulation by IL-4 and anti-inflammatory cytokines at the site of disease. Much of the increased expression in blood cells may be attributable to systemic release of TLR2 agonists from Mtb and can be mimicked with Pam3CysK4 and a macrophage line in vitro. A novel splice variant of TLR1, hsTLR1, was detected and showed the greatest response to Mtb in vivo and in vitro. Posttranscriptional stabilization of hsTLR1 and TLR1 mRNA after exposure to Mtb can account for increased expression in whole blood but is not sufficient to explain the increase in mRNA encoding hsTLR1 relative to that encoding TLR1.


   Acknowledgments
 
We thank Graham McIntyre for assistance in preparing and counting live organisms, Dr. Maria Manoussaka for kindly providing the cDNA of human B cells, and Dr. Martin Taylor (Wellcome Trust Centre for Human Genetics, Oxford, U.K.) for his bioinformatics assistance.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
G.A.W.R. is a research director and shareholder of SR Pharma PLC, which contributed to the laboratory costs of J.-S.C. However, SR Pharma PLC has no financial interest in the work or any relevant intellectual property rights.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This project was supported by a grant from SR Pharma. Back

2 Address correspondence and reprint requests to Dr. Graham A. W. Rook, Centre for Infectious Diseases and International Health, Royal Free and University College Medical School, 46 Cleveland Street, London W1T 4JF, U.K. E-mail address: g.rook{at}ucl.ac.uk Back

3 Abbreviations used in this paper: Mtb, Mycobacterium tuberculosis; ActD, actinomycin D; BAL, bronchoalveolar lavage; hsTLR1, human splice variant of TLR1; TB, tuberculosis; sMtb, sonicated Mtb; sMv, sonicated M. vaccae; UTR, untranslated region. Back

Received for publication September 13, 2005. Accepted for publication December 21, 2005.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

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