Evaluation of a commercial real-time PCR assay targeting Leishmania spp. with microscopically positive skin biopsies obtained from Nigerian patients

Abstract

Background

Although real-time polymerase chain reaction (PCR) assays for leishmaniasis are considered as highly sensitive, their standardization is still ongoing. In the study presented here, the sensitivity of a commercial real-time PCR assay targeting Leishmania spp. was evaluated with Nigerian skin biopsies from patients with microscopic diagnosis of cutaneous leishmaniasis (CL).

Methods

The samples were subjected to two subsequently applied types of nucleic acid extraction and afterwards characterized by two in-house real-time PCR assays targeting kinetoplast DNA (desoxyribonucleic acid) and the small subunit rRNA (ribosomal ribonucleic acid) gene of Leishmania spp. as well as by Sanger sequence analysis of the leishmanial hsp70 gene. Subsequently, the VIASURE Leishmania real-time PCR detection kit (CerTest Biotec, S.L., San Mateo de Gállego, Zaragoza, Spain) was applied with the DNA eluates.

Results

From 50 bioptic samples of Nigerian patients with CL, interpretable sequences could be obtained from 30 (60%), showing best matching with Leishmania major in all instances. The VIASURE assay confirmed the abundance of leishmanial DNA in all samples after both DNA extraction schemes, showing slightly better sensitivity compared to the reference PCR assays.

Conclusion

The VIASURE assay reliably detects L. major in microscopically positive bioptic samples from Nigerian CL patients.

1. Introduction

Leishmaniasis is an infectious disease caused by protozoan parasites and transmitted by sandflies [1]. During human infection, the protozoa replicate in macrophages [2]. Clinical manifestations range from localized skin infections over mutilating mucosal involvement to life-threatening systemic manifestations, depending on species, vector and immunological host factors [1, 2]. Saliva of the vector mosquitos is known to contain immunomodulatory agents affecting the manifestation of infections in the host [3–5]. Therapeutic interventions depend on manifestation and disease stage, comprising intralesional antimonial drugs for the treatment of skin efflorescences as well as systemic therapy with preferentially liposomal amphotericin B, miltefosine, antimonial drugs, pentamidine and azoles in case of systemic involvement [1]. Antimicrobial resistance has been described to limit the therapeutic success in a species-depending manner [1], while the status of adaptive immunity is a critical factor for successful therapy with relevance of both CD4+ and CD8+ T-lymphocytes [1, 6]. Vaccine development is in progress but so far, reliable vaccinations are not available [7].

Cutaneous leishmaniasis (CL), in particular, is prevalent in sub-tropical, tropical, and Mediterranean regions [8]. Potential regional shifts associated with the climate change have been recently evaluated as likely [9]. Although not life-threatening, scars resulting from leishmanial ulcers are feared because of the associated social stigma [10]. In fact, CL was recently confirmed as the third most frequent infectious skin disease after fungal infections and leprosy in refugees, migrants, asylum seekers, and internally displaced persons worldwide dwelling under over-crowded and resource-limited conditions with poor hygiene options [11].

While the usefulness of molecular diagnostics for the diagnosis of leishmaniasis is nowadays well accepted [10], comparably rare disease occurrence in resource-rich settings with associated low market shares for test producers makes certified commercial molecular diagnostic solutions still scarcely available. Light microscopy remains the standard diagnostic approach for CL in resource-limited settings, in spite of far-from-perfect diagnostic accuracy [2]. Approaches to increase the visibility of leishmania in microscopic samples, e.g., due to specific staining applying in-situ hybridization (ISH) or fluorescent in-situ hybridization (FISH) [12], have been described but did not lead to broad application, presumably because of the associated workload.

Leishmania-specific real-time PCR assays with excellent diagnostic accuracy in terms of sensitivity and specificity have been described for both genomic and kinetoplast DNA targets, but attempts on standardization and molecular quantification are still ongoing [13–15]. Comparative assessment of published real-time PCR protocols suggested kinetoplast minicircle DNA as the most promising molecular target in terms of diagnostic accuracy [15, 16], 18S rRNA-gene specific assays showed similarly reliable results [16]. While kinetoplast minicircle DNA-based assays usually show optimized sensitivity due to high sequence copy numbers in the target cells, they are poorly suited for the discrimination on subgenus or species level [17]. Notably, specificity estimates of Leishmania-specific PCR are usually better than reported sensitivity estimates, which is at least partly considered to be a consequence of interspecies target sequence variability [18]. Apart from first line diagnostic use, nucleic acid amplification testing (NAAT) was also found to be useful for confirmatory testing of microscopic test results [19]. For point-of-care application of Leishmania-specific NAAT, loop-mediated isothermal amplification (LAMP) was found to be a promising option, although its standardization is also yet ongoing [20]. PCR from skin smears showed comparable diagnostic sensitivity like PCR from aspirates or skin biopsies in a recent meta-analysis, making less invasively obtainable smear materials suitable for diagnostic testing [21]. Future technologies like combining NAAT with nano-biosensor technology or metagenomic screening approaches have been discussed [22, 23], but realistic implementation strategies still need to be convincingly demonstrated.

For a comparative test evaluation, skin biopsies from Nigerian individuals with microscopically diagnosed CL were used in the here-described study. Nigeria as well as its neighboring countries Cameroon and Chad are known for several decades as areas of endemicity of both cutaneous and visceral leishmaniasis [24–28], with phlebotomine vectors being abundant [28]. At the beginning of the 1990s, the regional prevalence of CL in Nigerian dermatology patients attending a university hospital was estimated in the 0.1% range [29]. Regional zoonotic transmission with small rodents like Mastomys natalensis and Tatera gambiana as animal hosts has been proposed for Nigeria [30]. Insofar, robust pre-test probability can be expected in Nigerian samples from patients with clinical suspicion of CL. The aim of the evaluation was a contribution to the so-far still scarcely available data on diagnostic accuracy of molecular testing for leishmania-specific DNA.

2. Methods

2.1. Study design and sample materials

The study was conducted as a head-to-head test comparison using highly sensitive real-time PCR assays targeting kinetoplast DNA (kDNA, later also referred to as reference PCR 1) and the small-subunit ribosomal RNA (ribonucleic acid) gene (SSU rRNA, later also referred to as reference PCR 2) of Leishmania spp. adapted from the literature [31, 32] as reference standards. The reference PCRs were augmented by a Sanger sequencing approach of the leishmanial hsp70 gene for confirmatory testing [33]. Residual sample materials of skin biopsies obtained from Nigerian individuals, which had been used for the microscopic diagnosis of leishmaniasis, were applied for the molecular assessments. Thereby, two different nucleic acid extraction approaches were compared as well. Patient-related data cannot be provided in line with ethical clearance for this methodical assessment as detailed below, which is an admitted deviation from the STARD criteria for diagnostic studies [34].

2.2. Molecular diagnostic approaches

Bioptic sample material from patients with the clinical suspicion of CL was stored at −80 °C. Nucleic acid extraction was initially conducted using the Chinese assay Nucleic Acid Extraction or Purification Kit (Magnetic Bead Method) (Hunan Runmie Gene Technology Co., Ltd, China; Code: RM-D-M1001) according to the manufacturer's instructions (later also referred to as nucleic acid extraction 1), including centrifugation, adsorption and filtration steps. As the initial nucleic acid extraction approach did not contain sample homogenization and protein digestion steps, a tissue protocol containing such steps was added to allow an in-depth assessment of the sample materials. The remaining sample pellet was subsequently subjected to additional nucleic acid extraction similar as described elsewhere [35] with some minor modifications. In short, pellets from all samples were manually mechanically homogenized using reaction tube pestles (Eppendorf, Hamburg, Germany) and 1.5 ml safe seal reaction tubes (Sarstedt, Nümbrecht, Germany). Afterwards, lysis with 12 units proteinase K (Qiagen, Hilden, Germany) for 5h or overnight at a temperature of 56 °C was conducted. Finally, nucleotide acid extraction from the lysed tissue homogenates was conducted using the commercial DNeasy Blood and Tissue kit (Qiagen, Hilden, Germany) as detailed in the manufacturer's protocol (later also referred to as nucleic acid extraction 2). Both types of DNA eluates were subjected to testing with the generic reference real-time PCRs targeting Leishmania spp.-specific kDNA [31] and the SSU rRNA gene [32] on RotorGene Q cyclers (Qiagen, Hilden, Germany) with minor adaptions as described [36]. Summarized, both assay were run in 20 µL volumes. The reaction chemistry of the kDNA assay [31] consisted of HotStarTaq Mastermix (Qiagen, Hilden, Germany), a final MgCl₂ concentration of 3.0 mM, 150 nM of each primer, 60 nM of the probe, and 2.0 µL sample DNA eluate. The run protocol comprised an initial denaturation step at 95 °C for 15 min. Afterwards, 45 cycles of denaturation for 15 s at 95 °C as well as of combined annealing and amplification for 60 s at 55 °C followed. For the SSU rRNA gene assay [32], HotStarTaq Mastermix (Qiagen, Hilden, Germany), a final MgCl₂ concentration of 2.0 mM, 200 nM of each primer, 20 nM of the probe, and 2.0 µL sample DNA eluate were used. Each run consisted of an initial denaturation step at 95 °C for 15 min, followed by 40 cycles of denaturation for 15 s at 95 °C as well as of combined annealing and amplification for 60 s at 60 °C. The applied oligonucleotides are shown in Table 1. Each PCR run included a positive control consisting of plasmids with target-specific sequence inserts (Table 1) and a PCR grade water-based negative control. Extraction and inhibition was controlled with a real-time PCR protocol targeting Phocid Herpes Virus (PhHV) DNA as previously described [37], which was performed by adding plasmids to the samples prior to the nucleic acid extraction procedure. There was not any specific cycle threshold (Ct) value cut-off for the definition of a positive real-time PCR signal. Each signal with a typical sigmoid shape was considered as a likely specific amplification.

Table 1.

Sequences of the oligonucleotides used for the leishmanial kDNA-specific reference PCR 1 [31], the SSU rRNA gene-specific reference PCR 2 [32], and for the sequence assessment of the leishmanial hsp70 gene [33], as well as insert sequences of the positive control plasmids for the kDNA PCR and the SSU rRNA gene PCR. Dashes were included in the shown oligonucleotide sequences to facilitate readability and do not delineate codons

Leishmania PCR according to Mary (reference PCR 1) [31]
Target gene	kDNA sequence
Forward primer Leish Mary F	5′-CTT-TTC-TGG-TCC-TCC-GGG-TAG-G-3′
Reverse primer Leish Mary R	5′-CCA-CCC-GGC-CCT-ATT-TTA-CAC-CAA-3′
Probe Leish Mary T, reporter and quencher	5′-FAM-TTT-TCG-CAG-AAC-GCC-CCT-ACC-CGC-BHQ1-3′
Sequence insert in the positive control plasmid	5′-CCA-CCC-GGC-CCT-ATT-TTA-CAC-CAA-CCC-CCA-GTT-TCC-CGC-CTC-GGA-CCC-GAT-TTT-TGA-CAT-TTT-TGG-CCA-ATT-TTT-GAA-CGG-GAT-TTC-TGC-ACC-CAT-TTT-TCG-ATT-TTC-GCA-GAA-CGC-CCC-TAC-CCG-GAG-GAC-CAG-AAA-AG-3′
GenBank accession number of insert sequence	EU437405.1
Leishmania PCR according to Wortmann (reference PCR 2) [32]
Target gene	SSU rRNA gene
Forward primer Leis U1 F	5′-AAG-TGC-TTT-CCC-ATC-GCA-ACT-3′
Reverse primer Leis L1 R	5′-GAC-GCA-CTA-AAC-CCC-TCC-AA-3′
Probe Leis P1 T, reporter and quencher	5′-FAM-CGG-TTC-GGT-GTG-TGG-CGC-C-BHQ1-3′
Sequence insert in the positive control plasmid	5′-AAG-TGC-TTT-CCC-ATC-GCA-ACC-TCG-GTT-CGG-TGT-GTG-GCG-CCT-TTG-AGG-GGT-TTA-GTG-CGT-C-3′
GenBank accession number of insert sequence	M81430.1
Sequencing PCR 1 (primers HSP70-F25 and HSP70-R617)* [33]
Target gene	hsp70
Forward primer HSP70-F25	5′-GGA-CGC-CGG-CAC-GAT-TKC-T-3′
Reverse primer HSP70-R617	5′-CGA-AGA-AGT-CCG-ATA-CGA-GGG-A-3′
Sequencing PCR 2 (primers HSP70-6F and HSP70-R1310)* [33]
Target gene	hsp70
Forward primer HSP70-6F	5′-GTG-CAC-GAC-GTG-GTG-CTG-GTG-3′
Reverse primer HSP70-R1310	5′-CCT-GGT-TGT-TGT-TCA-GCC-ACT-C-3′
Sequencing PCR 3 (primers HSP70-F25 and HSP70-R1310)* [33]
Target gene	hsp70
Forward primer HSP70-F25	5′-GGA-CGC-CGG-CAC-GAT-TKC-T-3′
Reverse primer R1310	5′-CCT-GGT-TGT-TGT-TCA-GCC-ACT-C-3′

*Amplification primers were used for Sanger sequencing as well.

In an attempt to confirm the results of leishmania-specific generic PCR and to further differentiate below the genus level, Sanger sequencing of the hsp70 gene of Leishmania spp. was conducted using a described protocol [33]. The applied oligonucleotides are shown in Table 1. Positive controls consisting of residual material from a positive sample and PCR grade water-based negative controls were included in each PCR run. Amplification of both the long reads and the two short reads was attempted [33]. Amplicons were visualized using the Lonza FlashGel system (Lonza Group, Basel, Switzerland). If visible bands were seen, the amplicons were transferred for nucleic acid extraction of the amplicons and subsequent Sanger sequencing of forward and reverse strands to the commercial company Microsynth Seqlab GmbH (Göttingen, Germany). Thereby, the amplification primers were used as sequencing primers as well. Forward and reverse strands were manually quality-controlled with the software Finch TV (Geospiza Inc., 2004–2012, Seattle, Washington, USA). Trimming and alignment of forward and reverse strands were conducted using the internet-based software DNA Subway 2.0 [38], nucleotide BLAST (basic local alignment search tool) assessment followed applying the nucleotide collection of NCBI (national center for biotechnology information) and restricting the search to Leishmania species. No sequence files were provided to international databases, because sequencing was merely diagnostically performed and not associated with sample-associated characteristics in line with the ethical clearance as detailed below. Aligned sequence reads are shown as Supplementary Material 1 together with this article.

The generic commercial assay VIASURE Leishmania real-time PCR detection kit (CerTest Biotec, S.L., San Mateo de Gállego, Zaragoza, Spain), which is cleared for diagnostic use in the European Union, was conducted on RotorGene Q cyclers (Qiagen, Hilden, Germany) as described by the manufacturer.

Applying a stronger and a weaker positive sample per assay, including eluates obtained with both nucleic acid extraction schemes, inter- and intra-assay variance was calculated based on three measurements each. For the intra-assay assessment, the sample was tested in triplicate in the same cycling run, while three different cycling runs were included in the inter-assay variance analysis.

2.3. Statistics

Raw data were arranged with the software Excel from the Office package of 2019 (Microsoft Corporation, Redmont, WA, USA). Sensitivity as compared to the diagnostic reference standard when assessed with the histologically positive sample materials including 95% confidence intervals (95% CI) including significance assessment based on Fisher's exact test as well as a non-parametric comparison of measured Ct-values of concordantly positive real-time PCR results using Wilcoxon matched pairs testing and of sequence-confirmed and not sequence-confirmed samples using Mann-Whitney-U testing were calculated applying the software GraphPad InStat, version 3.06 (GraphPad Software, Inc., San Diego, CA, USA). Also, this software was used to non-parametrically correlate Ct values obtained with the reference PCRs and the tested VIASURE assay using Spearman rank correlation testing. As the study was conducted without a priori hypotheses in an exploratory, hypothesis-forming manner, we abstained from correction for multiple-testing like, e.g., by applying strategies like the Bonferroni-Holmes correction [39], and considered each P-value <0.05 as significant with this simplified approach.

2.4. Ethics

All assessments were conducted in line with the Declaration of Helsinki and all its amendments. Ethical clearance was provided by the Medical Association of Hamburg, Germany (reference number: WF-011/19, obtained on 11 March 2019), allowing anonymous use of diagnostic residual sample materials for test comparison purposes without a need for informed consent.

3. Results

3.1. Sample characteristics

In 30 out of 50 assessed samples (60%), hsp70 gene sequencing showed best matching with deposited Leishmania major sequences. In all other samples, Sanger sequencing failed to provide interpretable sequence information. Details are indicated in the Supplementary Materials 1 and 2. Further, in all included samples, actual abundance of Leishmania spp. DNA was either confirmed by any of the applied reference real-time PCRs or the evaluated commercial real-time PCR assay. Details are shown in the Supplementary Material 3. Notably, sequence-confirmed samples showed lower Ct-values and thus higher quantities of target DNA than samples without sequence-based species assignment as L. major (Supplementary Material 4).

3.2. Assay sensitivity

When assessing the 50 microscopically positive bioptic samples, the VIASURE assay correctly identified all of them as positive, resulting in perfect diagnostic sensitivity after both nucleic acid extractions (Table 2). In contrast, the Leishmania kDNA-specific reference assay and the Leishmania SSU-rRNA-specific reference assay showed individual cases of failed identification following both nucleic acid extraction schemes (Table 2). Notably, the differences were not statistically significant, neither for the total number of assessed samples (Table 2) nor for the subpopulations of samples confirmed as L. major-positive by hsp70 gene sequencing (Supplementary Material 5). Inter- and intra-assay variance assessment is shown in Table 3.

Table 2.

Calculated sensitivity values of the assays. Possible differences between the different in-house assays and the commercial VIASURE assay over the two different modes of nucleic acid extraction were assessed applying Fisher's exact test

Assay	Nucleic acid extraction scheme	Calculated sensitivity (95% confidence interval)	Significance P applying Fisher's exact test
kDNA sequence assay	nucleic acid extraction 1	98.0% (89.4%, 100.0%)	P = 1.0000
	nucleic acid extraction 2	94.0% (83.4%, 98.7%)	P = 0.2424
SSU rRNA gene assay	nucleic acid extraction 1	98.0% (89.4%, 100.0%)	P = 1.0000
	nucleic acid extraction 2	96.0% (86.3%, 99.5%)	P = 0.4949
VIASURE assay	nucleic acid extraction 1	100.0% (92.9%, 100.0%)	Reference for nucleic acid extraction 1
	nucleic acid extraction 2	100.0% (92.9%, 100.0%)	Reference for nucleic acid extraction 2

Table 3.

Exemplary inter-assay and intra-assay variance assessment (3 recorded values each) over all applied real-time PCR assays and nucleic acid extraction schemes

Assay	Sample type	Nucleic extraction scheme	Inter-assay variation as variance of the recorded Ct values, in brackets: mean Ct value ±standard deviation	Intra-assay variation as variance of the recorded Ct values, in brackets: mean Ct value ±standard deviation
kDNA sequence assay	Stronger positive	2	0.09 (13.3 ± 0.3)	0 (13 ± 0)
kDNA sequence assay	Weaker positive	1	0.36 (27 ± 0.6)	0.09 (26.7 ± 0.3)
SSU rRNA gene assay	Stronger positive	1	0 (11 ± 0)	0 (11 ± 0)
SSU rRNA gene assay	Weaker positive	1	0 (30 ± 0)	0 (30 ± 0)
VIASURE assay	Stronger positive	2	0.09 (14.7 ± 0.3)	0.09 (14.3 ± 0.3)
VIASURE assay	Weaker positive	2	2.25 (34.7 ± 1.5)	0.09 (32.3 ± 0.3)

3.3. Influence of the nucleic acid extraction approach on the measured Ct values of the applied real-time PCR assays

The comparison of measured Ct values of the applied real-time PCR assays after both modes of nucleic acid reaction showed significant differences in three instances. First, a difference close to the significance threshold was observed for lower Ct-values after nucleic acid extraction 1 compared to nucleic acid extraction 2 for the assay targeting leishmanial kDNA. Also, significantly lower Ct-values after extraction scheme 1 were observed over the whole number of Leishmania spp.-specific real-time PCR results comprising all three tested assays. However, the dimension of recorded differences remained low in the 1-Ct-step-range and thus in a similar range as partly shown for the variance analyses in Table 3. In line with this, no significances were observed for the Leishmania SSU-rRNA-specific reference assay and the Leishmania spp.-specific reactions of the VIASURE assay, although there were non-significant tendencies for lower Ct values after extraction scheme 1 as well. In contrast, there was a robust significance in the opposite direction for the internal control reaction of the VIASURE assay, indicating substantial partial inhibition after nucleic acid extraction 1 compared to nucleic acid extraction 2. With an average difference of nearly 3 Ct-steps, this observation is in a different range than the intra- and inter-assay variance as shown in Table 3, with the exception of the inter-assay variance observed with the sample assessed with the VIASURE assay showing low quantities of target DNA. Details are provided in the Table 4.

Table 4.

Wilcoxon matched-pairs testing-based comparison of Ct values obtained with nucleic acid extraction 1 and 2

Assay	Mean Ct-value* after nucleic acid extraction 1 (± standard deviation)	Mean Ct-value* after nucleic acid extraction 2 (± standard deviation)	P as calculated applying Wilcoxon matched-pairs testing
kDNA sequence assay	24.4 (±5.3)	25.8 (±4.7)	0.0342
SSU rRNA gene assay	22.5 (±4.7)	23.4 (±4.4)	0.1656
VIASURE assay	24.6 (±5.0)	25.2 (±4.8)	0.0919
Fused assessment of Ct-values obtained with all three Leishmania spp.-specific assays	23.9 (±4.9)#	24.8 (±4.7)#	0.0027
Internal control PCR of the VIASURE assay	23.3 (±1.0)	20.7 (±0.7)	<0.0001

*Only samples showing positive results with both compared nucleic acid extraction procedures were included. P = significance level. ^#Numeric values by themselves should not be interpreted, because they were generated with different assays.

3.4. Correlation of the Ct values measured applying the Leishmania-spp. reference real-time PCR assays with the results of the Leishmania spp.-specific reactions of the VIASURE assay

While absolute numbers of measured Ct values varied in an assay-dependent manner, there was imperfect yet significant correlation between Ct values measured with the Leishmania spp. reference real-time PCR assays and the results of the Leishmania spp.-specific reaction of the VIASURE assay as nonparametrically indicated applying Spearman rank correlation testing. Notably, higher Spearman r values were recorded after nucleic acid extraction 1 compared to nucleic acid extraction 2. Details are shown in Table 5.

Table 5.

Nonparametric correlation as indicated by Spearman rank correlation testing when comparing Ct-values of the in-house assays with Ct-values of the VIASURE assay for concordantly positive samples

Assay comparison	Nucleic extraction assay	Spearman r	95% confidence interval	Significance P
kDNA vs. VIASURE	1	0.4937	(0.2386, 0.6850)	0.0003
kDNA vs. VIASURE	2	0.3890	(0.1060, 0.6138)	0.0069
SSU rRNA gene vs. VIASURE	1	0.4386	(0.1713, 0.6459)	0.0016
SSU rRNA gene vs. VIASURE	2	0.3241	(0.03537, 0.5630)	0.0246

4. Discussion

The study was conducted to evaluate the sensitivity of a commercial Leishmania spp.-specific PCR assay with DNA from 50 skin biopsies of Nigerian individuals with CL against two reference real-time PCRs and Sanger sequence-based confirmation of the abundance of Leishmania spp.-specific DNA. In a previous study with samples from patients with CL from Pakistan applying the same assay, 100% sensitivity had been demonstrated [40], while sensitivity of 81.8% was shown with a sample panel from patients with either CL or visceral leishmaniasis [41] and sensitivity of 95% with infected sand flies [42]. Two modes of nucleic acid extraction, which were applied subsequently with the same sample materials due to lacking sample homogenization and protein digestion in the course of the first extraction scheme, were included in the assessment. The investigation led to a number of results.

First of all, the assessed commercial real-time PCR assays showed similarly high sensitivity, even without failed amplification in all assessed samples, compared to the reference real-time PCRs, which had been identified as highly sensitive in previous assessments [31, 32, 36]. While the commercial assay identified all 50 microscopically positive samples after both nucleic acid extraction schemes, individual missed samples were found for both competitor assays and modes of nucleic acid extraction.

The traditional block-cycler-based sequencing PCR targeting the hsp70 gene of Leishmania spp. turned out to be less sensitive than the applied real-time PCR assays. Accordingly, sequence-based confirmation succeeded in 60% only with subsequent Sanger sequencing. Thereby, all obtained sequences showed best matches with L. major, which is known to be abundant in west Africa [43].

In direct comparison of both applied nucleic acid extraction procedures, Leishmania spp.-specific Ct values were measured in similar ranges with a tendency for slightly lower Ct values after nucleic acid extraction 1. In comparison, the samples showed considerably higher rates of partial sample inhibition after nucleic extraction 1 as indicated by higher Ct values in the internal control PCR. Considering the partial inhibition after extraction 1, it must be concluded that this mode of extraction resulted in a higher yield of target DNA, compensating for this partial inhibition and leading to similar Ct values after both extraction schemes. A likely explanation is reduced abundance in the sample pellets, which were subjected to nucleic acid extraction 2 after initial nucleic acid extraction 1. As the recorded Leishmania spp.-specific Ct values were nevertheless in a similar range, it is likely that nucleic acid extraction scheme 2 might have resulted in even lower Ct values if it had been performed with fresh sample materials rather than with pre-extracted pellets. Unfortunately, lacking availability of more sample materials did not make a more direct comparison feasible. Nevertheless, the findings underline the statement in the manufacturer's instruction of the VIASURE assay that the assay can be applied with “any commercially available DNA extraction kit“ in line with the respective manufacturer's instructions. Notably, this does not only include the assessed manual Qiagen extraction assay but also the company's solution for highly automated nucleic acid extraction applying the EZ1 Advanced automated instrument (Qiagen, Hilden, Germany) [44].

The producers of the assessed commercial VIASURE assay did not include details on the target sequence in their test manual, however, substantial correlation with the results of reference real-time PCR assays targeting kDNA and the SSU rRNA gene of Leishmania spp. could be shown. This correlation indirectly confirms that the different applied real-time PCR assays most likely targeted a common meta-structure, although Sanger sequence-based identification of L. major succeeded in a proportion of samples only.

The assessment has a number of limitations. First of all, the selective inclusion of bioptic samples from patients with CL limits the interpretation to sensitivity only, while statements cannot be given on the assay's specificity. The reason is the focus of the analysis on the assay's performance with this specific sample type and study population, not on a comprehensive repetition of the manufacturer's validation in the course of the assay's in-vitro diagnostics accreditation. Second, the limited number of available samples limits the assessment's interpretability, resulting in broad 95% confidence intervals and the associated risk of overlooking minor differences. For the same reason, nucleic acid extraction 2 could only be conducted from pellets remaining after initial application of nucleic acid extraction 1. Third, the ethical clearance allowing the use of the samples for diagnostic accuracy assessment purposes did not allow providing sample-related clinical information, limiting the interpretation to a sole technical comparison.

5. Conclusions

In spite of the abovementioned limitations, the assessed Leishmania spp. DNA-specific VIASURE real-time PCR showed 100% sensitivity with bioptic samples taken from patients with lesions microscopically assigned to the diagnosis CL in Nigerian individuals. In terms of not missing any included microscopically positive samples, the commercial PCR assay performed even slightly better than in-house competitor assays which have shown similar diagnostic accuracy estimates in previous assessments [31, 32, 36].

Supplementary materials

eujmi-16-231-s001.pdf ^{(406.7KB, pdf)}

Abbreviations

The following abbreviations are used in this manuscript:

95% CI

95% confidence interval

BLAST

basic local alignment search tool

CL

cutaneous leihmaniasis

Ct

cycle threshold

DAT

direct agglutination test

DNA

desoxyribonucleic acid

FISH

fluorescent in situ hybridization

HIV

human immunodeficiency virus

ISH

in situ hybridization

kDNA

kinetoplast DNA

LAMP

loop-mediated isothermal amplification

n.a.

not applicable

NAAT

nucleic acid amplification technique

NCBI

national center for biotechnology information

PCR

polymerase chain reaction

PhHV

Phocid herpes virus

rk39

39-amino acid repeat within recombinant kinesin

rRNA

ribosomal ribonucleic acid

SSU

small subunit

Funding Statement

Funding: External funding was not received for this investigator-initiated assessment.