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Isoenzyme analysis of cultured parasites is the conventional approach for Leishmania species identification. Molecular approaches have the potential to be more sensitive and rapid. We designed PCR generic primers to amplify a segment of the rRNA internal transcribed spacer 2 (ITS2) from multiple Leishmania species. To validate the selected ITS2 fragment, we tested clinical specimens and compared the species results obtained by the molecular approach (PCR followed by DNA sequencing analysis) with those from the parasitologic approach (in vitro culture followed by isoenzyme analysis). Among the 159 patients with clinical specimens positive by both approaches, a total of eight Leishmania species were identified. The species results were concordant for all but two patients: for one patient, the results were Leishmania (Viannia) guyanensis by the molecular approach versus L. (V.) braziliensis by the parasitologic approach; for the other patient, the results were L. (Leishmania) tropica versus L. (L.) major, respectively. ITS2 PCR, followed by sequencing analysis, can be used to detect and discriminate among Leishmania species. The results confirmed our hypothesis that a region of the ITS2 gene can complement the characterization of Leishmania parasites at the species level. The approach we developed can be used as a diagnostic tool in reference laboratories with adequate infrastructure to perform molecular characterization of pathogens.
Leishmaniasis encompasses multiple clinical syndromes, most notably, visceral, cutaneous, and mucosal forms (9, 13, 14, 22, 27, 30). Leishmaniasis is endemic in focal areas in the tropics, subtropics, and southern Europe and is caused by species in the Leishmania and Viannia subgenera (14). Different species can be associated with diverse clinical manifestations and sequelae. Species identification can facilitate clinical management, such as decisions regarding whether/which treatment is indicated.
Isoenzyme analysis of cultured parasites is the conventional diagnostic approach for Leishmania species identification. Molecular approaches have the potential to be more sensitive and rapid; e.g., the results can be available within days versus weeks or months. However, Leishmania PCR-based assays have varied in sensitivity, specificity, and utility for genus- or complex-level versus species-level characterization (2, 3, 7, 10, 24–26, 28); some approaches have been highly sensitive for genus-level detection, e.g., PCR methods that target minicircle kinetoplast DNA (kDNA), whereas others have been tailored for particular species/settings.
We sought to develop a comprehensive molecular approach for characterizing Leishmania species; we focused on the rRNA internal transcribed spacer (ITS) region, which has been sequenced by other investigators and proposed as a target for molecular typing (12, 25). Here we describe our identification and validation of a diagnostic fragment that was amplified from the ITS2 with a PCR generic primer pair that we designed. Rather than conduct a formal evaluation of the molecular approach, we focused on demonstrating proof of concept. By using ITS2 PCR, followed by DNA sequencing analysis of the amplicons, we were able to discriminate among Leishmania spp. to complement the diagnosis of leishmaniasis. This method has been incorporated into the Centers for Disease Control and Prevention (CDC) algorithm for the laboratory diagnosis of leishmaniasis, improving the turnaround time for diagnosis and for decisions regarding treatment of cases of infection.
For the initial evaluations of the Leishmania ITS2—to identify what we refer to as the diagnostic fragment—we analyzed a total of 40 cultured isolates previously characterized by isoenzyme analysis and epidemiologic criteria as Leishmania (Viannia) braziliensis, L. (V.) guyanensis, L. (V.) panamensis, L. (L.) mexicana, L. (L.) amazonensis, L. (L.) donovani, L. (L.) infantum, L. (L.) chagasi, L. (L.) aethiopica, L. (L.) major, and L. (L.) tropica (Table 1).
To validate the diagnostic ITS2 fragment, we analyzed clinical specimens that were submitted from 2005 to 2009 to our reference laboratory at the CDC for Leishmania testing. Specimens were used in accordance with a CDC human subject-approved protocol; no additional specimens were obtained for the molecular analysis. Per routine, to the extent possible, the specimens were shipped by overnight mail; almost all of the specimens were from skin biopsies, and specimens typically were shipped in a sterile tube containing RPMI medium that CDC provided in advance. Upon arrival at the CDC, specimens were divided into portions for the parasitologic and molecular analyses described below. When possible, stained slides (e.g., histopathology slides provided by the practitioner or impression smears/wet mounts made by the CDC) were examined by light microscopy for amastigotes; the slides were not necessarily from the same specimens that were tested by culture and PCR.
The culture tubes contained Novy-MacNeal-Nicolle agar with 10% defibrinated rabbit blood and an overlay of RPMI medium with 15% fetal bovine serum. The tubes were incubated at 25°C; aliquots were examined by light microscopy two to three times a week until promastigotes were noted or 4 weeks had elapsed. Positive cultures were transferred from tubes to large flasks containing ~20 ml of fresh RPMI medium to facilitate obtaining large numbers of parasites for isoenzyme analysis. In brief, after the parasites again reached log phase, the liquid culture medium that was harvested from the flasks was centrifuged at 850 × g for 15 min (in phosphate-buffered saline [PBS], pH 7.2), the pellet was frozen, and fresh RPMI medium was added to the flasks, which were placed back into the incubator. This process was repeated several times, using the same tube to harvest the pellet. After the final harvest, the composite pellet was treated with 50 to 100 μl of lysis buffer (0.1 M Tris, 0.1 M malic acid, 0.1 M EDTA, and 0.1 M MgCl2); the mixture was subjected to several freeze-thaw cycles at −70°C, and the lysate was stored in liquid nitrogen pending isoenzyme analysis. Cellulose acetate electrophoresis was performed, based on protocols developed by Kreutzer et al. (15–17), targeting isoenzymes 6-phospho-d-gluconate dehydrogenase (6PGDH) and glucose phosphate isomerase (GPI).
DNA was extracted from the 40 banked, frozen cultured isolates—also referred to as the reference isolates—by using a QIAamp DNA Micro Kit (Qiagen), following the manufacturer's instructions. To amplify the ITS region, we used a previously described strategy (12) and cloned the fragments as described below. To identify/select the diagnostic ITS2 fragment, we conducted multiple alignments of the full-length ITS sequence (data not shown); the sequence of the ITS2 region of each Leishmania species was identified by searching the GenBank database using the BLAST algorithm. On the basis of this analysis, we found a variable region within nucleotide positions 392 and 809 [using L. (L.) donovani GenBank entry FJ753386 as a reference], designed the generic primers LGITSF2 (GCATGCCATATTCTCAGTGTC) and LGITSR2 (GGCCAACGCGAAGTTGAATTC) for this region, and used this primer pair to reamplify the DNA previously extracted from the 40 reference isolates.
For PCR, the total volume per reaction mixture was 50 μl, which included 1 μl of DNA, 0.2 μM each primer, 0.2 mM each deoxynucleoside triphosphate (dNTP), 1.5 mM MgCl2, and 2.5 U of AmpliTaq Gold PCR polymerase (Applied Biosystems). PCR was performed in a GeneAmp PCR System 9700 thermocycler (Applied Biosytems), using the following cycle structure: 95°C for 5 min, followed by 40 cycles of 95°C for 30 s, 60°C for 30 s, and 72°C for 1 min, with a final extension of 72°C for 7 min. The amplicons were purified with a StrataPrep PCR Purification Kit (Stratagene) and cloned with a pCR2.1-TOPO vector TOP10 Kit (Invitrogen). The clones were purified by using a QIAprep Spin Miniprep Kit (Qiagen) and sequenced by using BigDye, version 3.1, chemistry (Applied Biosystems). The sequencing reaction mixtures were purified by using MultiScreen-HV plates (Millipore) and were analyzed on an ABI Prism 3100 sequencer, with data collection software, version 2.0, and DNA Sequencing Analysis Software, version 5.1 (Applied Biosystems). The sequences were assembled, edited, and aligned by using the DNA STAR SeqMan (DNASTAR Inc.) and the GeneStudio suite (GeneStudio Inc.) programs.
We also designed an alternative reverse generic primer named LGITS2R1 (GAATTCTCGTTTTGGTTTTTTG), at positions 773 to 794 of the L. (L.) donovani reference (GenBank entry FJ753386), to compensate for the CG deletion and the G/T transversion in the LGITS2R2 primer binding region of L. (L.) amazonensis and L. (L.) aethiopica, respectively. We compared the PCR results obtained with three primer combinations—LGITSF2/LGITSR2, LGITSF2/LGITSR1, and LGITSF2/LGITSR1/LGITSR2—on seven cultured isolates of L. (L.) amazonensis and three of L. (L.) aethiopica, besides at least five isolates of each of the other pertinent species (Table 1). The amplicons were resolved with 1.5% agarose gels and visualized by staining with ethidium bromide.
DNA was extracted directly from the clinical specimens (i.e., from ~5 to 10 mg of tissue and ~100 to 200 μl of blood or bone marrow) by using the method described above. PCR amplifications using generic primers LGITSF2/LGITSR2 were performed in a final reaction volume of 50 μl as described above, except that the aliquot of specimen DNA was 5 (versus 1) μl, diluted at 1:1 and 1:5. Sequencing reactions were performed directly from the amplicons by using the parameters described above.
The initial molecular testing of the clinical specimens was blinded such that isoenzyme results were not known. However, for some specimens, if the species results differed from those obtained by the parasitologic approach (e.g., negative versus positive), the molecular testing was repeated. If residual specimen was available, DNA was reextracted; otherwise, DNA from the original extraction was reamplified. The species results obtained by isoenzyme analysis were considered the gold standard.
Each DNA extraction batch included a negative control, i.e., DNA extracted from Leishmania-free eukaryotic cell cultures (E-6 cells in RPMI medium). For each PCR run, a tube containing water and PCR mix was used as a negative control (blank), and approximately 1 pg of the cloned full-length ITS region was used as a positive control. Separate rooms were used for DNA extraction, PCR Master Mix preparation, DNA loading onto the Master Mix, loading of the positive control, and electrophoretic analysis of amplicons.
Use of the ITS2 generic primer pair LGITSF2/LGITSR2 to amplify DNA from the 40 cultured isolates produced amplicons that ranged from 372 to 450 bp in length (Table 1). The PCR results obtained by using different primer combinations—LGITSF2/LGITSR1, LGITSF2/LGITSR1/LGITSR2, or LGITSF2/LGITSR2—were comparable (Fig. 1 A, B, and C), which suggests that either or both reverse primers (LGITSR1 and LGITSR2) can be used with the forward primer LGITSF2. We used LGITSF2/LGITSR2 to accomplish the validation described below.
Alignment of the ITS2 sequences showed that amplicons from the same species or even the same specimen varied in length, mainly because of AT, TC, TG, or GA microsatellite repeats (Table 1). To simplify the analysis, these discrepant repeat regions were removed, which yielded intraspecies sequences of comparable lengths. However, because of a few transitions and transversions, more than one sequence per species or isolate was found. Therefore, the few variant sequences were removed, and the resulting identical sequences were considered the consensus sequences.
In some instances, direct sequencing analysis of LGITSF2/LGITSR2 amplicons showed degraded regions, with multiple overlapping peaks after well-resolved regions, because of the regions with variant repeats. One example is the AT microsatellite starting at nucleotide positions 118 and 129 observed in the amplicons obtained from L. (V.) braziliensis (MHOM/BR/75/M2903) and L. (L.) tropica (MHOM/SU/74/K27) WHO reference strains. Although the partial sequences sufficed for correct species identification by BLAST search, we cloned the reference strain amplicons to obtain the complete sequence of the fragment. Alignment of sequences obtained by randomly selecting one clone per species (from the identical group of sequences) showed sufficient divergence for discrimination both between and among species complexes/subgenera.
The L. (L.) donovani species complex had a stretch of Gs (a G stretch) that started at nucleotide position 343 [using L. (L.) donovani FJ753386 as a reference]. The reference strains of L. (L.) infantum and L. (L.) chagasi were indistinguishable; therefore, we have used the nomenclature “L. (L.) infantum/chagasi.” However, the G stretch typically was longer for L. (L.) infantum/chagasi (7 or 9 Gs) than for L. (L.) donovani (5 Gs). Two of the four L. (L.) donovani strains, including the WHO strain from Asia, had a G/T transversion at position 345, referred to as G345T (Fig. 2).
Among the sequences for the three Viannia spp. in the analyses (Table 1), we consistently found sites for species-level identification in the clones that we sequenced from each isolate; the species were differentiated on the basis of four single nucleotide polymorphisms (SNPs), i.e., an A/C transversion at nucleotide position 59 (referred to as A59C transversion), a G220A transition, a G297A transition, and a G317A transition [positions are based on the sequences of L. (V.) braziliensis FJ753376, L. (V.) guyanensis FJ753387, and L. (V.) panamensis FJ948438]. Leishmania (V.) braziliensis was distinguished from the other Viannia spp. by the presence of nucleotides A, G, G, and G at positions 59, 220, 297, and 317, respectively; L. (V.) guyanensis and L. (V.) panamensis were distinguished from each other on the basis of the transition G220A. Figure 2 shows the differences that allowed species identification.
The ITS2 fragment was amplified directly from clinical specimens (versus cultures), the amplicons were sequenced, and the species was identified on the basis of criteria described for the reference isolates, including the SNPs and G stretch (Fig. 2). If direct sequencing analysis showed overlapping peaks because of variant microsatellites, partial sequences were used to identify the species; no cloning was done.
Species-level characterization was accomplished by this molecular approach (PCR followed by DNA sequencing analysis) for specimens from 202 patients, including 13 (6%) for whom the positive results were obtained on repeat testing (Table 2). We compared the species data with data from the parasitologic approach (in vitro culture followed by isoenzyme analysis). Of necessity, the comparison was limited to the 159 patients for whom species results were obtained by both approaches. For all but 2 of these 159 patients, the results were concordant. One of the two exceptions was in the Viannia subgenus: L. (V.) guyanensis was identified by the molecular approach, whereas L. (V.) braziliensis was identified by the parasitologic approach. DNA from this specimen was reamplified and resequenced thrice, and the same result was obtained each time. The other exception was a discordance between L. (L.) tropica (by PCR and sequence analysis) and L. (L.) major (by culture and isoenzyme analysis). DNA from these specimens was reamplified and resequenced twice, and the same results were obtained each time.
The comparative data were limited to the L. (L.) donovani species complex as a whole because isoenzyme analysis using 6PGDH and GPI does not distinguish L. (L.) donovani from L. (L.) infantum/chagasi. In general, the molecular approach appeared to be more sensitive than the parasitologic approach for detecting parasites in this species complex; i.e., specimens were positive for 29 versus 14 patients, respectively. By using molecular criteria described for the reference isolates (Fig. 2), L. (L.) infantum/chagasi was the etiologic agent for all but 6 of these 29 patients, including 14 of the 15 with negative cultures. For the other six patients, the species was identified by the molecular approach as L. (L.) donovani; four of these six patients became infected in Central and South America, where L. (L.) infantum/chagasi would be the expected species (Table 2).
Various molecular approaches for diagnosing Leishmania infection have been described. For example, PCR methods that target minicircle kDNA can be highly sensitive for detecting Leishmania parasites because of the abundance of minicircles in each kinetoplast (2, 7, 10). However, high-level sequence polymorphism among minicircles is an impediment for species identification with protocols based solely on kDNA PCR. Targeting the rRNA-coding cluster poses other issues (7, 32). Highly sensitive approaches for identifying particular Leishmania species (or a species complex/subgenus) have been described (8, 25). However, if multiple Leishmania spp. need to be differentiated in a diagnostic laboratory, molecular approaches that require different PCR primers for each species have the potential for carryover contamination. Although strategies to minimize this potential have been developed (5), contamination has been observed even when strict protocols were followed (20, 23).
Our goal was to develop a molecular approach for species-level discrimination that requires only one pair of PCR primers. We focused on the ITS region: the rRNA internal spacers are subject to less evolutionary pressure and show more sequence divergence than the coding regions and have been proposed as targets for molecular typing (1, 6, 19). We identified a region of the ITS2 adequate for our diagnostic goal and designed generic PCR primers (LGITSF2/LGITSR2) to amplify this fragment from Leishmania spp. associated with human infection. We found substantial differences in the ITS2 region spanned by these primers that generally allowed species identification.
During our analysis of the reference strains, we noted intraspecies variability in the AT, TC, TG, or GA microsatellite repeats, as have other investigators (12, 18, 25, 31), which initially hindered use of the ITS2 fragment for species identification. We also noted some distinct sequences because of a few transitions and transversions. In most instances, distinct sequences were found in clones obtained from the same isolate, which likely indicated the presence of multiple different ITS copies in the same genome. To exclude the possibility of erroneous incorporation of nucleotides (11, 29), we reamplified the DNA from some specimens by using Pfu High-Fidelity DNA polymerase (Stratagene); the results were the same as those with the routinely used Taq DNA polymerase (data not shown). During the development phase of the assay, we cloned reference strain amplicons to obtain the sequence of the entire diagnostic fragment for each species, and we demonstrated that partial sequences sufficed for species identification. Therefore, in the validation phase, when we tested clinical specimens, we used direct sequencing analysis, without cloning, and in some instances relied on partial sequence data.
The distribution of species among the clinical specimens [e.g., the disproportionate number of patients infected with L. (V.) panamensis] reflects travel and immigration patterns among U.S. civilians evaluated for leishmaniasis; almost all of the clinical specimens we analyzed were from skin lesions. Although the PCR results obtained with different combinations of the reverse primers LGITSR1 and LGITSR2 were comparable for the isolates tested (Fig. 1A, B, and C), we had insufficient numbers of specimens positive for L. (L.) amazonensis and L. (L.) aethiopica—the species that prompted us to design the alternative reverse primer (LGITS2R1)—to compare the sensitivities of the primer pairs. We correctly identified the species as L. (L.) amazonensis for seven cultured isolates (including two WHO reference strains). However, this species was not identified by the parasitologic or molecular approach for any clinical specimens (Table 2). For L. (L.) aethiopica, the analyses included three cultured isolates (including one from WHO) and two clinical specimens (Tables 1 and and22).
The overall agreement between the molecular and parasitologic approaches was >98% when the comparison, of necessity, was limited to specimens for which a species determination was accomplished by both approaches. The two discordant results (among 159 patients) remained unresolved: for one patient, L. (V.) guyanensis was identified by the molecular approach versus L. (V.) braziliensis by the parasitologic approach (culture followed by isoenzyme analysis), and for the other patient, L. (L.) tropica was identified versus L. (L.) major, respectively. No other methods were used to evaluate the specimens; both results were epidemiologically plausible (Table 2), and the possibility of coinfection or of variant strains could not be excluded.
In general, our molecular approach differentiated among Viannia spp., namely, L. (V.) braziliensis, L. (V.) guyanensis, and L. (V.) panamensis, on the basis of four SNPs that appear to be in linkage disequilibrium with the genes that code for the isoenzymes we used to differentiate them (i.e., 6PGDH and GPI). These species, each of which likely is heterogeneous, have been associated with phenotypic (clinical) differences, which underscores the importance of species-level diagnostic techniques (4, 14, 22).
Our molecular approach did not differentiate L. (L.) infantum from L. (L.) chagasi, which many experts consider synonymous (18, 21). However, some distinguishing features in the ITS2 fragment were noted between the reference isolates of L. (L.) donovani versus L. (L.) infantum/chagasi, i.e., the length of a G stretch and, for the WHO L. (L.) donovani strain from Asia, a G435T transversion. Similar variation has been reported previously for some L. (L.) donovani strains (18). However, for several clinical specimens [four of the six classified as L. (L.) donovani], the approach we used appears to have resulted in misclassification of L. (L.) infantum/chagasi as L. (L.) donovani. Additional experience with more specimens from more areas of endemicity is needed to ascertain the ability of this ITS2 molecular target to reliably distinguish L. (L.) donovani from L. (L.) infantum/chagasi.
Approximately half of the patients with negative cultures but positive PCR results had slides available that were positive by light microscopic examination (Table 2), thereby confirming the diagnosis of leishmaniasis and underscoring the plausibility of PCR positivity. In some instances, negative culture (versus molecular) results might have been attributable to fastidious or nonviable parasites in the specimens we received or to contamination of the cultures, e.g., because of suboptimal handling or delayed shipment to the CDC. Lack of PCR positivity for some of the culture-positive specimens might have been attributable to heterogeneous distributions of parasites in specimens, such that the parasite/DNA concentration in the portion analyzed by PCR was below the detection limit of the assay. Preliminary data from a 40-cycle SYBR green-based real-time PCR assay under development in our laboratory suggest that positive specimens with threshold cycle (CT) values around 35 could be negative by ITS2 PCR and/or culture (M. E. de Almeida, unpublished data).
A drawback of our approach is the need for DNA sequencing analysis, which is not yet available in most laboratories in areas of endemicity. However, the ITS2 fragment has cleavage sites for HinP1I or HhaI enzymes (GCGC) and MnlI enzyme (GAGG) that may have utility for identifying the species we evaluated except for those in the Viannia subgenus. A cleavage site for HphI (GGTGAN7) could be used to identify L. (V.) guyanensis but not L. (V.) braziliensis or L. (V.) panamensis. These cleavage sites could be explored as targets for restriction fragment length polymorphism analysis to distinguish among a few species by laboratories in which DNA sequencing analysis is not yet available.
In summary, our findings confirm our hypothesis that a region of the ITS2 gene can complement the characterization of Leishmania parasites at the species level. The approach we developed can be used as a diagnostic tool in reference laboratories that have adequate infrastructure to perform molecular characterization of pathogens.
We are indebted to Patricia Wilkins from the Centers for Disease Control and Prevention for invaluable assistance in reviewing the manuscript.
The findings and conclusions in this report are those of the authors and do not necessarily represent the official position of CDC.
Published ahead of print on 13 July 2011.
†The authors have paid a fee to allow immediate free access to this article.