INTRODUCTION

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Leprosy continues to be a significant health problem globally. In 1995, the World Health Organization reported that the number of registered leprosy patients was 1.3 million, while the estimated number was closer to 1.8 million (27). Although multidrug therapy has been very successful in reducing the prevalence of the disease, the annual incidence has not yet declined in most countries where the disease is highly endemic. Furthermore, a significant number of patients with leprosy have nerve damage and disabilities at the time of diagnosis. Although it has become clear in recent years that subclinical infection is quite common, the epidemiology of leprosy is still poorly understood. Reliable methods for the identification of subclinically infected individuals or other potential reservoirs for the spread of the disease and methods for the early detection of patients with leprosy before disability occurs are not yet available.

There is no "gold standard" for the diagnosis of leprosy. The disease is generally diagnosed on the basis of clinical criteria. As in many other centers, slit skin smears stained to detect acid-fast bacilli (AFB) are used to confirm the diagnosis and classification in the All Africa Leprosy Rehabilitation and Training Center (ALERT) hospital and leprosy control program in Ethiopia. For patients with diagnostically difficult cases of infection, skin or nerve biopsy specimens are obtained and diagnosis is made on the basis of characteristic histological findings and the presence of AFB within the biopsy specimen. Because acid-fast staining requires at least 10⁴ organisms per gram of tissue for reliable detection (4), sensitivity is low, particularly for patients at the tuberculoid end of the leprosy spectrum when AFB are rare or absent. However, microscopy is used because Mycobacterium leprae cannot be cultivated in vitro and immunological antigen or antibody detection methods are too insensitive. Recently, a number of investigators have used PCR to amplify various genomic sequences of M. leprae in order to improve detection when low numbers of bacteria are present (1, 5, 6, 8, 11, 15, 23, 24, 28).

In this study, we have developed an alternative detection method which targets the abundant 16S rRNA of M. leprae. Detection of rRNA should impart increased sensitivity over assays based on the detection of a single copy or even multiple copies of genomic sequences since each cell contains 1,000 to 10,000 copies of rRNA. An RNA-based detection method would be expected to better reflect the number of viable organisms because RNA is generally degraded within a few minutes of cell death. Thus, an RNA-based detection system might be useful for confirmation of the diagnosis in patients for whom a diagnosis is difficult to make, for assessing the efficacy of chemotherapy, in distinguishing relapse from late reaction in previously treated patients, and for epidemiological studies.

In developing our reverse transcription (RT)-PCR assay, we have chosen primers which span regions of the 16S rRNA-coding sequence unique to M. leprae in order to ensure species specificity. We have tested both the species specificity and the sensitivity of our assay. Furthermore, we have demonstrated its sensitivity and specificity in detecting M. leprae in tissue biopsy specimens.

	MATERIALS AND METHODS

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Patient samples. Skin biopsy specimens (4-mm punch) were obtained from newly diagnosed, untreated leprosy patients seen at the ALERT hospital in Addis Ababa, Ethiopia, after obtaining informed consent. Twenty-one of these patients were classified clinically as having paucibacillary (PB) leprosy (20 borderline tuberculoid and 1 borderline lepromatous) and 29 were classified clinically as having multibacillary (MB) leprosy (9 polar lepromatous, 20 borderline lepromatous). Skin samples were bisected, and half of each sample was fixed in buffered formalin for subsequent hematoxylin and eosin or acid-fast staining, while the other half was mounted with cryoembedding medium, flash frozen, and stored at 80°C for RT-PCR. Biopsy specimens were histologically classified according to the scale of Ridley and Jopling (16).

RNA isolation. Forty cryostat sections 5 µm thick were cut from frozen biopsy specimens by using a fresh blade for each sample. The biopsy specimens were placed in a guanidinium isothiocyanate-based RNA isolation buffer (RNA STAT-60; Tel-Test, Friendswood, Tex.) while they were still frozen, homogenized with 0.1-mm-diameter glass beads, and then sonicated for 5 min at 60°C in a water bath (Transsonic, Elma, Germany) at a frequency of 35 kHz. The remainder of the RNA isolation (phenol-chloroform extraction and isopropanol precipitation) was performed according to the manufacturer's instructions.

cDNA synthesis. RNA (2 µg) was transcribed into cDNA by using avian myeloblastosis virus reverse transcriptase (Stratagene, La Jolla, Calif.) in a 20-µl reaction volume containing 50 mM Tris-HCl (pH 8.5), 8 mM MgCl₂, 30 mM KCl, 6 mM dithiothreitol, a 0.25 mM concentration of each deoxynucleoside triphosphate, 1 nM synthetic oligo(dT)15, 1 nM random hexamers, 1 nM P3 primer, and 400 U of RNase inhibitor (Stratagene) at 42°C for 50 min. The mixture was then heated to inactivate the enzymes, cooled, and diluted to 100 µl with sterile RNase-free distilled H₂O before it was added to the PCR mixture. For experiments to determine whether transcribed RNA or contaminating DNA was amplified by our protocol, purified nucleic acid was subjected to RNase or DNase treatment prior to cDNA synthesis, as described by Huang et al. (9). The cDNA reaction mixture was prepared as usual, except that 500 ng of RNase A or RNase-free DNase (Boehringer Mannheim) was added to the reaction mixture in place of avian myeloblastosis virus reverse transcriptase. The RNase inhibitor was omitted in reactions containing RNase. The mixture was incubated for 30 min at 37°C, followed by incubation for 5 min at 75°C to inactivate the enzyme. Reverse transcriptase was then added directly to the mixture and cDNA synthesis was completed by our standard method.

Oligonucleotides. All oligonucleotides were purchased from the DNA analysis facility at Johns Hopkins University School of Medicine in Baltimore, Md. The sequences of primers P1 and P3 were originally published by Arnoldi et al. (1), and the sequence of primer P2 was originally published by Cox et al (5). The primers and their sequences were as follows: GAPDH upstream (positions 367 to 386, mRNA), ACC ACC ATG GAG AAG GCT GG; GAPDH downstream (positions 875 to 894, mRNA), GTG GAA GGA CTC ATG ACC ACA GTC CAT GCC; GAPDH probe (positions 571 to 600, mRNA), CTC AGT GTA GCC CAG GAT GC; M. leprae 16S rRNA P1 (positions 9 to 28, DNA), AGA GTT TGA TCC TGG CTC AG; M. leprae 16S rRNA P2 (positions 69 to 91, DNA), CGG AAA GGT CTC TAA AAA ATC TT; M. leprae 16S rRNA P3 (positions 218 to 239, DNA), CAT CCT GCA CCG CAA AAA GCT T; and M. leprae 16S probe (positions 86 to 112, DNA), CGC CAC TCG AGT ATC TCT AAA AAA GATT.

PCR. PCR was performed in a total volume of 50 µl of buffer obtained from Boehringer Mannheim (10 mM Tris HCl [pH 8.3], 1.5 mM MgCl₂, 50 mM KCl) or Stratagene (10 mM Tris-HCl, 1.5 mM MgCl₂, 75 mM KCl [pH 8.3]) containing 5 mM deoxynucleoside triphosphates, a 1 µM concentration of each of the upstream and downstream primers, 5 to 10 µl of template (cDNA or genomic DNA), and 2.5 U of Taq DNA polymerase (Boehringer Mannheim) with a Hybaid Omnigene thermal cycler. Negative controls, which contained all reaction components except template, were included in all experiments to detect contamination. The cycling profile for the two sets of 16S rRNA primers involved 40 cycles of denaturation at 94°C for 2 min, annealing at 60°C for 2 min, and extension at 72° for 3 min for 40 cycles, followed by final extension at 72°C for 5 min. For amplification of GAPDH, the samples were heated to 94°C for 3 min initially and were then kept at 94°C for 1 min, followed by annealing at 60°C for 1 min and extension at 72°C for 2 min for 30 cycles, with an additional final extension time of 5 min.

Agarose gel electrophoresis and Southern blotting. Aliquots (15 µl) of the PCR products were electrophoresed through 2% agarose gels. DNA was transferred to Hybond N⁺ nylon membranes (Amersham) after depurination in 0.25 N HCl for 15 min and denaturation in 0.5 N NaOH for 30 min with a vacuum blotter (Bio-Rad) for 90 min at a vacuum pressure of 5 in. Hg, or overnight by capillary transfer. DNA was fixed to the nylon by cross-linking upon exposure to 150 mJ of UV light (Bio-Rad). After prehybridization for 30 min at 47°C, the blot was hybridized to a fluorescein-labeled oligonucleotide probe (10 ng/ml) prepared according to the manufacturer's instructions (ECL 3'-oligo labeling and detection system; Amersham, Little Chalfont, United Kingdom) at 47°C for 2 to 17 h. Membranes were washed sequentially in 5× SSC (1× SSC is 0.15 M NaCl plus 0.015 M sodium citrate) with 0.1% sodium dodecyl sulfate twice for 5 min at room temperature and then in 1× SSC with 0.1% sodium dodecyl sulfate twice for 15 min at 47°C. Chemiluminescence detection was performed according to the manufacturer's instructions. Briefly, the membranes were incubated with horseradish peroxidase-conjugated anti-fluorescein antibody which catalyzes the generation of light upon exposure to luminol in the detection solution. Light was detected by exposure of Hyperfilm-ECL (Amersham) to membranes.

Isolation and enumeration of M. leprae for sensitivity testing. Isolation of mycobacteria from the patients' biopsy specimens was performed at the ALERT hospital clinical laboratory by the method of Rees (26). Fresh biopsy specimens were homogenized in 0.1% bovine serum albumin with a Pyrex homogenizer. A known volume of tissue suspension was serially diluted and was spread with a calibrated loop over 8-mm-diameter circles scored on clean microscope slides, dried, fixed, and stained by the modified Ziehl-Neelsen method. The number of bacteria in eight oil-immersion fields (magnification, ×1,000) were counted, and the original concentration was calculated. The morphologic index (MI), which was the percentage of solid-staining bacilli (22), was determined after examining 100 bacilli lying separately.

Determination of sensitivity. To assess the sensitivity of the method with isolated and counted M. leprae, serial 10-fold dilutions of the bacterial suspension were prepared in 50 µl of phosphate-buffered saline (1.9 mM NaH₂PO₄, 8.1 mM Na₂HPO₄, 154 mM NaCl [pH 7.2]) before solubilization in RNA STAT-60. Samples were processed as described above, except that 1 µl of glycogen (20 mg/ml) was added to each tube during the precipitation step to enhance the yields.

Bacterial strains. The sources and strain numbers of the bacterial species used to test the specificity of the assay are listed in Table 1. All mycobacteria except M. leprae were grown in Lowenstein-Jensen medium at 37°C. The Rhodococcus strains were grown in Sabouraud dextrose medium at 30°C, and both Propionibacterium and Corynebacterium strains were grown aerobically and anaerobically at 37°C on both blood and chocolate agars. Staphylococcus aureus and Streptococcus pneumoniae were grown on blood agar.

	RESULTS

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We tested several sets of primers for their efficiency of detection (data not shown). Primers P1 and P3 reproducibly amplified a 231-bp fragment when M. leprae cDNA prepared from RNA purified with commercially available reagents was used, while primers P2 and P3 amplified a 171-bp fragment under the same conditions. We then showed that the 171-bp product is amplified from transcribed RNA and not from small amounts of genomic DNA contaminating our RNA preparations by pretreating identical samples with either RNase or DNase prior to cDNA synthesis and PCR. RNase treatment prevented amplification of the 171-bp product, while DNase treatment had no effect, as indicated in Fig. 1. In addition, no PCR product was obtained when isolated RNA was subjected directly to PCR (data not shown).

View larger version (38K): [in this window] [in a new window]   FIG. 1.   RNAse and DNase treatment of isolated nucleic acid. Replicate RNA samples were subjected to nuclease treatment prior to cDNA synthesis, PCR for M. leprae with primers P2 and P3, electrophoresis, and hybridization of an internal probe to Southern blots of the 171-bp PCR products. Lane 1, no nuclease treatment, lane 2, RNase; lane 3, DNase; lane 4, negative control; lane 5, additional positive control RNA.

To assess species specificity, we purified RNA from M. leprae, 28 other mycobacterial species, seven related genera, and three organisms which are commonly found on the skin or in the noses of healthy individuals. After cDNA synthesis, we subjected each sample to amplification using primer set P1-P3 and primer set P2-P3. The results obtained by probing Southern blots of the RT-PCR products are summarized in Table 1. Primer set P2-P3 amplified a 171-bp product only when M. leprae cDNA served as the template. In contrast, a 231-bp product was amplified when primer set P1-P3 was used in the PCR of cDNA from all mycobacterial species tested, but this product was not amplified from the cDNA of closely related nonmycobacterial species or organisms normally found on the skin and in the nose (Table 1). Thus, we concluded that primer set P2-P3 is species specific, while primer set P1-P3 is likely to be genus specific.

We next assessed the sensitivity of the assay. M. leprae was isolated from fresh skin biopsy specimens from untreated lepromatous leprosy patients who were seen at the ALERT hospital or from lymph nodes of a nude mouse which had been infected with M. leprae since the bacteria cannot be cultured in vitro. The isolated bacteria were counted microscopically, and the MI was determined after acid-fast staining. RNA was extracted from serially diluted organisms, followed by cDNA synthesis, PCR, electrophoresis, and Southern blot analysis. The results of a typical experiment with organisms isolated from tissue from a nude mouse are presented in Fig. 2. Amplified cDNA was readily visible when 10 organisms were present. When the experiment was repeated with organisms derived from the skin lesion of a lepromatous leprosy patient, 23 organisms were detected (data not shown). In each case the MI, which is related to viability (12), was less than 10%. The assay therefore appears to be quite sensitive.

View larger version (14K): [in this window] [in a new window]   FIG. 2.   Sensitivity of RT-PCR for M. leprae. Serial tenfold dilutions of counted M. leprae were made in 50 µl of phosphate-buffered saline. Each tube was subjected to RNA extraction, RT, PCR, electrophoresis, Southern blotting, hybridization, and detection by enhanced chemiluminescence analysis. Lane 1, 106 bacteria; lane 2, 105 bacteria; lane 3, 104 bacteria; Lane 4, 103 bacteria; lane 5, 102 bacteria; lane 6, 10 bacteria; lane 7, 1 bacterium; lane 8, 101 bacteria; lane 9, 102 bacteria; lane 10, 103 bacteria; lane 14, positive control.

We then tested the ability of the assay to detect M. leprae within skin biopsy specimens from 50 untreated leprosy patients seen at the ALERT hospital. RT-PCR for GAPDH served as a positive control for each sample, as illustrated in Fig. 3. M. leprae was detected in 82% of the skin biopsy specimens from newly diagnosed leprosy patients (Table 2). This included 32 of 33 (97%) biopsy specimens which contained AFB on microscopic examination of at least six sections and 9 of 17 (53%) skin biopsy specimens which were negative for AFB. Only one bacterium was seen on careful examination of the single specimen that was negative by RT-PCR but positive for AFB. The assay was positive for 96% (25 of 26) of the biopsy specimens from patients with MB disease and 67% (12 of 18) of the patients with PB disease who were not thought to be undergoing a reaction on clinical grounds. Patients thought to be undergoing a reaction on clinical grounds were less likely to have a positive RT-PCR test result (one of five patients overall; none of two patients with borderline tuberculoid disease and a clinical reaction, none of two patients with borderline lepromatous disease and a clinical reaction, and one patient with polar lepromatous disease and a clinical reaction). The RT-PCR assay detected M. Leprae in more biopsies than acid-fast staining of either slit skin smears or biopsy specimens (Table 2). No false-positive results were obtained; RT-PCR of skin biopsy specimens obtained from patients with a variety of other inflammatory skin diseases and healthy individuals gave negative results (Table 2).

View larger version (28K): [in this window] [in a new window]   FIG. 3.   Detection of M. leprae in skin biopsy specimens from leprosy patients. Each biopsy specimen was subjected to RT-PCR, agarose gel electrophoresis, Southern blotting, hybridization with a fluorescein isothiocyanate-labeled probe, and detection by enhanced chemiluminescence analysis. Row 1, PCR with GAPDH primers; row 2, PCR with M. leprae primers. Lanes 1 to 14, aliquots of the RT-PCR product amplified from skin biopsy specimens from different patients; lane 15, negative control; lane 16, positive control.

	DISCUSSION

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We have shown that M. leprae can be detected by RT-PCR of the 16S rRNA. We found that use of one of the upstream primers (primer P2) described by Cox et al. (5) in combination with the oligonucleotide used as a probe by Arnoldi et al. (1) conferred species specificity in our PCR assay. The primer of Cox et al. (5) anneals to an AT-rich sequence within the 12-bp insertion that is found in the first variable region of the 16S rRNA gene. This insertion is unique to the M. leprae 16S rRNA genomic sequence compared to the sequences of other bacterial species (5, 19). In contrast to the assays based on 16S rRNA sequences developed by other investigators (1, 21), demonstration of species specificity by our assay does not require hybridization to a probe, although the sensitivity of the RT-PCR is improved by probe hybridization to Southern blots of PCR products (data not shown). The fact that our detection system does not require radioactive probes is an additional benefit.

Our assay has a theoretical advantage in terms of its greater sensitivity over those of protocols that detect a single copy or even multiple copies of genes since each bacterium has many copies of 16S rRNA. As a member of the slowly growing group (5, 17, 19) of mycobacteria, M. leprae has one copy of the 16S rRNA gene (17) but an estimated 4,000 molecules of 16S rRNA per cell (7). This estimate may be low, since the viability of M. leprae isolated from tissue is generally poor and the presence of a high proportion of dead bacteria in the sample used may have artificially decreased the estimate of the number of rRNA molecules per cell.

We were able to detect as few as 10 organisms from infected mouse tissue and 23 organisms from human tissue, even though the MI for each sample was less than 10%. Other investigators have obtained similar estimates of sensitivity (1 to 100 organisms) by performing PCR with serially diluted nucleic acid. However, we extracted RNA after the organisms isolated from infected tissues were serially diluted and counted microscopically since the exact amount of 16S rRNA within each cell is unknown. This method is subject to the inherent error involved in using a bacterial loop to make smears and in estimating the viability of M. leprae on the basis of the MI, which requires skilled staff and precise staining conditions.

Other investigators (23) have indicated that PCR inhibitors were present in some of their skin samples, particularly those with heavy infiltration. Inhibitors are less likely to influence our assay because our procedure includes extraction and precipitation of RNA rather than immediate PCR amplification of crude cell lysates. Although we did not rigorously test for inhibitors, they did not appear to be a major problem in our studies because we were able to amplify the control gene, GAPDH, from all but one of our samples with only 30 amplification cycles. The one GAPDH-negative biopsy specimen was unexpectedly faintly positive by the M. leprae RT-PCR assay. This finding may have been due to poor preservation of the RNA in this sample, the presence of inhibitors, or the high relative copy number of the M. leprae 16S rRNA in this borderline lepromatous biopsy specimen with a BI of 5.

We have validated the utility of our assay in detecting M. leprae in clinical specimens. The sensitivity of detection of M. leprae in skin biopsy specimens by RT-PCR was similar to the sensitivity of PCR methods that detect DNA (6, 28). Although one would predict that our assay would be more sensitive than those based on DNA targets, overall sensitivity may be decreased if the RT-PCR assay detects only viable organisms, in contrast to assays based on DNA detection. One would also predict that all specimens positive for AFB would be positive by PCR. We found one specimen that was positive for AFB but RT-PCR negative. This biopsy specimen contained only one acid-fast bacillus on careful examination of six sections. Since the numbers of bacteria were so low, it is possible that no organisms were present within the tissue aliquot that was processed for RT-PCR. Alternatively, the scarce bacteria within this specimen may have been nonviable since this biopsy specimen was classified histologically as borderline tuberculoid.

Detection of RNA rather than DNA may be useful for the detection of viable organisms. The half-life of mRNA may be as little as 2 min following cell death. Several investigators have shown that mRNA detection is a reliable indicator of cell viability. Patel et al. (14) showed that heat treatment of M. leprae reversed their ability to detect M. leprae by an RT-PCR assay that detected a 71-kDa heat shock protein, while Bej et al. (3) showed that the viability of Legionella was related to the levels of mRNA for the macrophage infectivity potentiator protein. However, the survival time of rRNA is less certain. The fact that ribosomes rapidly disappear when mycobacterial cells are degraded (18) suggests that the rRNA found within these structures might also be degraded shortly after cell demise. Recently, van der Vleit et al. (20) demonstrated a strong correlation between the isothermic RNA amplification product of the 16S rRNA of Mycobacterium smegmatis and the numbers of CFU after treatment of cultures with various doses of rifampin and ofloxacin. In contrast, DNA amplification of genomic DNA encoding the 16S rRNA gene did not correlate with the viability assessed by in vitro culture.

Our RT-PCR assay may also indicate the presence of viable organisms. We showed that our protocol detects rRNA and does not seem to be affected by the small amounts of genomic DNA that can contaminate some RNA preparations isolated by the guanidinium isothiocyanate method. To ensure that no DNA is amplified when one wishes to measure viability, one could include a DNase treatment step prior to cDNA synthesis for all samples, as described by Huang et al. (9).

The RT-PCR assay may be useful in a number of clinical situations. In addition to aiding in the diagnosis of difficult cases of PB leprosy and facilitating epidemiological studies, it may be useful for assessing the response to chemotherapy. Jamil et al. (11) applied limiting dilution PCR of the DNA encoding the 36-kDa pra antigen for five patients undergoing multidrug therapy to assess the response to chemotherapy. They reported a correlation coefficient of 0.5 between the number of organisms detected by PCR and the MI for the biopsy specimens. Although this is an encouraging result, one would postulate that a better correlation between the number of organisms detected by RT-PCR and the MI or some other measure of viability would be found. An assay that detects viable organisms would be useful for the determination of whether persisters, i.e., those organisms present at the completion of two or more years of multidrug therapy (2, 25), are viable and therefore are a potential source of relapse, especially in patients with MB leprosy (10). Similarly, the assay might be useful in distinguishing a relapse from a late reaction.