FormalPara Key Points

Due to the limited number of currently available anti-leishmanial drugs, effective clinical management, chemotherapy, and control of transmission depend largely on early and unequivocal diagnosis. Any patient residing in a visceral leishmaniasis (VL) endemic area presenting with a history of fever of more than 2 weeks’ duration and with no response to antibiotics/antimalarials should be tested for VL with an rK-39 dipstick test.

The rK39 rapid diagnostic test (RDT), used within strict clinical criteria, is currently used with good results for diagnosis of VL, but this test cannot differentiate between active disease and past VL. Furthermore, a diagnostic algorithm of the rK39 RDT for detection of asymptomatic infection has only been validated for high-incidence settings in strict combination with clinical criteria (fever for more than 2 weeks’ duration plus an enlarged spleen).

With the advent of technology, highly specific and sensitive molecular-based tools have been developed for detecting infection, diagnosis, and species differentiation and hold considerable further promise for delivering better point-of-care diagnostic tests in the elimination and post-elimination setting.

Molecular-based methods play a key role in early diagnosis, monitoring of treatment effectiveness, and assessment of drug resistance in Leishmania parasites.

1 Introduction

Leishmaniasis has been identified as high-priority disease by the World Health Organization (WHO) [1]. It is caused by protozoa belonging to the genus Leishmania and transmitted by the bite of a 2–3 mm long insect vector, the phlebotomine sand fly found throughout the world’s inter-tropical and temperate regions [2]. Around 21 species of Leishmania are known to be pathogenic to humans [3].The disease occurs in three forms: self-healing or chronic cutaneous leishmaniasis (CL), mutilating mucosal or muco-cutaneous leishmaniasis (ML or MCL), and life-threatening visceral leishmaniasis (VL). Each form varies in degree of severity, with VL being by far the most devastating with the highest mortality.

VL, also known as kala-azar, the most severe form of leishmaniasis, is caused by the obligate intracellular protozoan parasites Leishmania donovani/L. infantum. It is estimated by the WHO that 200,000–400,000 new cases of VL occur annually worldwide, 90% of which occur in three geographical regions: (1) South-East Asia—India (especially Bihar), Bangladesh, and Nepal; (2) Latin America—mainly north-eastern Brazil; and (3) East Africa—Sudan, Ethiopia, Kenya, Uganda, and Somalia [4,5,6,7]. In the Indian subcontinent (ISC), VL is now being reported in 54 districts in India, 16 upazila (administrative regions) in Bangladesh, and 12 districts in Nepal [8]. In Europe, VL is endemic in nine countries, accounting for less than 2% of the global burden [9]. In Brazil, VL is endemic in 21 of 26 states and a total of 14,859 cases were reported between 2001 and 2014 in 25% of Brazilian municipalities [10]. However, in Africa-VL is endemic in 17 localities in seven states in Sudan [11] and 6 regional states in Ethiopia.

The Leishmania parasite is transmitted by female Phlebotomine sand flies as a flagellated, metacyclic promastigote, which is phagocytized by host macrophages and then differentiates into the non-flagellated, replicative amastigotes [12]. The organs commonly affected during VL are the bone marrow, liver, and spleen [12]. Thus, clinical symptoms include hepatosplenomegaly, which is characterized by an enlarged abdomen with a palpable spleen and liver. Other symptoms include long-term, low-grade fever, muscle wasting, anemia, leukopenia, polyclonal hyper-gammaglobulinemia, and weight loss [13, 14]. If left untreated, it has a mortality rate of almost 100%. During an epidemic in the early 1990s in Sudan, there were an estimated 100,000 deaths. Risk of an epidemic still exists in the horn of Africa, at the junction of Eritrea, Ethiopia and Sudan, a highly endemic region where tens of thousands of refugees, returnees, and agricultural workers have been resettled. Especially in Sudan, Ethiopia, Kenya, Uganda, and Somalia, VL is the cause of much morbidity and mortality, and only a small minority of patients have access to diagnosis and treatment [15]. VL is endemic in several tropical and subtropical regions and has been reported in 56 countries around the world (Fig. 1). Importantly, the disease affects mostly poverty-stricken people, with over 80% of patients living below the poverty threshold (daily income of less than US$1) whose source of income is agriculture and/or animal husbandry [16]. More than 75% live in mud or grass-covered houses. These patients are thus completely dependent on charity or public health services for diagnosis or treatment, and these remain grossly deficient in endemic areas [17, 18].

Fig. 1
figure 1

Map of the global distribution of visceral leishmaniasis

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Importantly, the three countries affected by VL on the Indian subcontinent, India, Nepal and Bangladesh, aspired to eliminate VL by 2015 (a deadline later reset to 2020). The aim is to reduce the incidence to less than 1 per 10,000 of population at the sub-districts level (i.e., block level in India and Nepal and upazila level in Bangladesh) through early diagnosis, complete treatment of cases, and integrated vector management [19]. However, as countries move towards elimination goals, the number of VL and post-kala-azar dermal leishmaniasis (PKDL) (characterized by skin lesions in which parasites can be identified, in a patient who is otherwise fully recovered from VL) cases will decrease, and a low number of such cases will almost inevitably lead to a decreasing awareness in the communities and by health providers. If cases of VL and PKDL are ignored or missed in such a context, a new epidemic phase may start. To avoid such a scenario, there is need for development and validation of an innovative set of tools to find cases of VL and PKDL, along with an outbreak management strategy, and surveillance methods for the measurement of infection. Thus, there is a direct need for new technology  for the monitoring of infections, treatment effectiveness, and drug resistance because such validated methods under routine conditions do not exist. Molecular detection tools would constitute a more rapid and high-throughput alternative to detect parasites. In this review, we discuss the various molecular methods, focusing on recent developments and their clinical application in Leishmania detection, absolute quantification, species differentiation, and phylogenetic analysis.

2 Standard Diagnostic Tools and Their Limitations

A major challenge in the clinical management of VL is the weakness of health systems at the primary health centre (PHC) level in many affected countries, with multiple challenges and numerous constraints [8, 20]. Despite multiple techniques for confirming VL cases being available, they are all still far from being ideal. To date, observation of parasites in splenic aspirate is considered the gold standard for VL diagnosis [21]. While microscopic examination of spleen aspirates is a rapid and cheap approach with high sensitivity and specificity, it is not practical at the PHC level. However, in Kenya, VL policy specifies that all serologically proven leishmaniasis be confirmed by spleen aspirate, a procedure that can only be performed in referral hospitals [22]. Alternative parasitological diagnostic techniques are the lymph node or bone marrow aspirates (reviewed in Singh and Sundar [23]), the standard means of diagnosing VL in most countries. Sensitivity of such diagnostic methods is highly variable and dependent on the sampling procedure and technical skills of the physician or personnel performing the tests. Although sensitivity of bone marrow is lower than splenic aspiration, the diagnostic potential in combination with serology is adequate for clinical purposes [24, 25]. Again, these methods cannot be performed at the PHC setting in endemic areas because they require skilled personnel, have a high cost, and are less simple. Most importantly, examination of bone marrow/splenic aspirates is now only recommended when the rK39 rapid diagnostic test (RDT) is negative but the suspicion of VL disease is high or in VL patients diagnosed by rK39 who do not respond to first-line treatment [23]. Notably, very few practitioners currently have the skills to perform these dangerous aspirates and very limited numbers of these biopsies are taken in the Indian subcontinent. Culturing Leishmania promastigotes from tissue biopsies/peripheral blood mononuclear cell/whole blood is another method of diagnosis but is expensive and requires a sophisticated laboratory [26].

Human VL is associated with high level of plasma antibodies; however, although it is useful in diagnosis, the role of antibodies in VL pathogenesis is not clear [27, 28]. A number of non-invasive serological tests to detect Leishmania antibodies are now available. The Direct Agglutination Test (DAT) has been extensively validated in endemic areas and is recommended by the WHO for VL control programs [21, 29]; however, the requirements of relatively specific material and expertise make its use difficult in peripheral health centers. Similarly, the Indirect Immunofluorescent Antibody Test (IFAT) requires an immune-fluorescence microscope, which restricts its use to referral hospitals. Hope is now directed at a rapid immunochromatographic test based on a recombinant 39-amino acid repeat antigen (rK39 dipstick), which, despite the variability initially observed among different producers and countries, seems to be the first choice for decentralized diagnosis of VL and has good sensitivity and specificity [30, 31]; however, it shows decreased sensitivity in East Africa when compared with the Indian subcontinent [31, 32]. The rK39 dipstick is stable, easy to use, and is a ‘rapid test’ (results available in 10 min). Performance of this test has been comprehensively reviewed in various studies recently [23, 24, 31]. In India, Bangladesh, and Nepal, the VL elimination initiative has adopted the rK39 RDT as its main tool, but it has significant limitations as it cannot be used to diagnose relapses or to assess response to treatment (test of cure) [23, 33]. Approximately 10–20% of healthy people living in endemic areas test positive with the rK39 RDT [34, 35]. Importantly, despite its limitations, the rK39 RDT has been, and is still, a great asset in the struggle against VL as it allows diagnosis and treatment to be decentralized to as close as possible to the villages where patients live. However, although the rK39 RDT test represents a sound approach in highly endemic areas, this is not the case in situations of low infection intensity. Furthermore, these antibody detection tests are of limited used in immunocompromised patients (i.e., HIV co-infection) [23]. Antigen detection is required as a means of identifying symptomatic infections in immunocompetent and immunocompromised patients (e.g., diagnosis of primary VL in Sudan where rK39 RDTs lack sensitivity and complex diagnosis of VL relapse is required) and as an indicator of cure. The latex agglutination test (KAtex; Kalon Biological, Guildford, UK) detects the leishmanial antigen in urine and records the results in a scoring system that correlates well with the parasite load. However, KAtex is currently not considered to be an ideal test as it had poor sensitivity when tested at different centers [32, 36,37,38].

2.1 Molecular Diagnosis and Detection of Infection

Most Leishmania species have been sequenced, revealing an overall conservation of gene order, chromosome structure, and discrete differences in gene content. These recent research advancements have helped in the development of more appropriate rapid molecular diagnostic devices and platforms [39]. However, despite the technological development, there is a huge difference in using a commercially available and standardized molecular diagnostic as opposed to in-house kits. So far, several molecular methods have been developed for detection, identification, quantification, and phylogenetic analysis, and these are summarized in Fig. 2.

Fig. 2
figure 2

Molecular tools and markers for diagnosis of visceral leishmaniasis. A2 amastigote stage gene, cpb cysteine protease B, EF1 elongation factor-1, ELISA enzyme-linked immunosorbent assay, G6PD glucose-6-phosphate dehydrogenase, HSP heat-shock protein, hsp70 heat-shock protein 70, kDNA kinetoplast DNA, LAMP loop-mediated isothermal amplification, MALDI-TOF Matrix assisted laser desorption ionization - time of flight, MLEE multilocus enzyme electrophoresis, MLMT multilocus microsatellite typing, NGS next generation sequencing, NASBA nucleic acid sequence-based amplification, OligoC oligochromatography test, PCR polymerase chain reaction, polA DNA polymerase α, RAPD random amplified polymorphic DNA, SNP single nucleotide polymorphism

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The development of polymerase chain reaction (PCR) kits has provided one of the most sensitive and specific methods for diagnosis of clinical VL; they amplify parasite DNA and can be visually read without sophisticated equipment [44, 78,79,80]. The sensitivity of the PCR assay mostly depends on the biological sample (e.g., blood, bone marrow, splenic fluids, etc.) and the primers used to amplify the target sequence (variable or conserved target region) [81, 82]. The most commonly used amplification targets are nuclear DNA such as the small subunit ribosomal RNA (SSU rRNA) gene [45, 83, 84], extra-chromosomal DNA such as repetitive kinetoplastid DNA (kDNA) [45, 46, 85], mini-exon genes [86], and the ribosomal internal transcribed spacer (ITS) region [52]. A comparative overview of frequently used PCR targets and its sensitivity and specificities in different tissue samples are summarized in Table 1. One of the major limitation of DNA based PCR is the counting dead parasite DNA (as half life of DNA is 24 h within the body, which is still controversial and not proven); thus, RNA based amplification target is preferred [82, 87]. However, reliable RNA extraction is difficult in PHC settings. Srivastava et al. [44] validated 18S rRNA-based PCR in the blood of the largest number of patients and controls in their study, and found a sensitivity of 87.8% (95% confidence interval [CI] 84.1–89.8) and specificity of 94.6% (95% CI 92.8–96.1). Leishmanial DNA has been detected by PCR in the peripheral blood of persons with asymptomatic infection in Brazil and this was also documented recently in India and Nepal [88,89,90]. Several cohort studies conducted in India, Nepal, Bangladesh, Italy, Ethiopia, Sudan, and Brazil for detection of asymptomatic L. donovani infection in endemic villages has confirmed the increased capacity of PCR tests to detect infection in healthy individuals [89,90,91,92,93,94]. PCR assays have also been performed using non-invasive samples, such as buccal swabs and urine, with sensitivity of 79–83 and 88–97%, respectively [95, 96]. Molecular diagnosis using PCR is very useful in HIV–VL patients in whom the clinical picture is confusing and serological as well as immunological tests are not reliable due to low sensitivity [97]. Furthermore, sensitivity and specificity of PCR for detection of low-level parasitemia have been shown to be improved significantly by performing nested and semi-nested PCR, which involves two sets of primers (targeting a single gene locus) used in two successive runs. The second set of primers amplify the secondary target within the product of the first PCR product [49] but nested PCRs are prone to contamination and are not recommended except in accredited laboratories. The sensitivity and specificity of nested PCR using SSU-rRNA in the diagnosis of VL are reported to be 97% and 100%, respectively [98]. Similarly, multiplex PCR involves amplification of different DNA targets at the same time [99]. Although such assays are more sensitive than conventional PCR, their high costs make this test inappropriate in a field setting. Other forms of PCR such as the OligoC-TesT (Coris BioConcept, Gemblous, Belgium) [100], PCR-ELISA (enzyme-linked immunosorbent assay) [101], and nucleic acid sequence-based amplification (NASBA) have been developed and found to be more sensitive than conventional PCR [43]. More recently, rapid and highly specific loop-mediated isothermal amplification (LAMP) has emerged as a powerful tool for point-of-care diagnosis and has been validated in VL and PKDL in several countries [102, 103]. One of the advantages of this assay is that the test can be performed without the need for sophisticated equipment, making it a more attractive tool for field-based diagnosis. This assay is more rapid and cost effective than conventional PCR, but is limited in utility due to false positivity. Importantly, PCR-oligochromatography and LAMP are the only tests available commercially and this offers huge benefits over in-house kits in terms of reliability—of course, this comes at a price. Most recently, a recombinase polymerase amplification (RPA) assay (a simple and molecular assay as a mobile suitcase laboratory) was developed for canine VL [104]. This assay has also been tested and proved a promising diagnostic method for VL, which would significantly decrease the cost associated with testing [105].

Table 1 Comparison of molecular methods for detection, differentiation, identification, and quantification of Leishmania species
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Importantly, in view of mounting drug pressure, PCR diagnostic assays play a key role in monitoring drug efficacy and early reporting of drug resistance, which are essential to bring corrective actions in drug policy; this is even more important when the drug arsenal is limited, as in the case of VL [122,123,124]. The molecular assays are the only standard, rapid, high-throughput, and easy methods to track parasite resistance that can completely replace tedious in vitro susceptibility assays. Furthermore, such tools should be as simple as possible to be applicable and affordable in endemic countries. Recently, Srivastava et al. [125] identified a single nucleotide polymorphism in the cysteine proteinase B (cpb) gene that is associated with amphotericin B drug resistance.

2.2 Quantification of Parasites (Severity of Disease)

As an analytical technique, the conventional PCR method has some limitations. By first amplifying the DNA sequence and then analyzing the product, quantification is exceedingly difficult as the PCR gives rise to essentially the same amount of product independent of the initial amount of DNA template molecules that were present. Therefore, a conventional PCR (qualitative analysis) test shows only the presence or absence of Leishmania without quantification of the parasite load. With the highly efficient detection chemistry, sensitive instrumentation, and optimized assays that are available today in real-time PCR (also known as quantitative real-time PCR if DNA is the starting genetic material for quantification of parasites), the number of DNA molecules of a particular sequence in a complex sample can be determined with unprecedented accuracy and sensitivity that is sufficient to detect a single molecule. However, quantitative PCR (qPCR) does not directly measure the number of viable parasites circulating in the blood, but rather the amount of circulating parasite DNA. Therefore, sensitivity of qPCR depends on the assay design (primer and target region), chemistry used (SYBER® Green [Sigma-Aldrich, St Louis, MO, USA] or TaqMan® [Life Technologies, Foster City, CA, USA]), nature of clinical samples (blood, skin, bone marrow, or splenic fluids), and the DNA extraction methods (manual vs. commercial kits) [126] (Table 2). Using this technique, it was earlier demonstrated that the simultaneous quantitative evaluation of Leishmania DNA and cytokines by real-time PCR assay allows prediction of the development of disease in asymptomatic infected dogs [127]. Using qPCR, we have shown that the parasite load decreases during treatment in VL cases. Amplification of the 18Sr RNA gene sequence from a small volume of heparinized whole blood using real-time PCR revealed a wide range of blood parasitemia in VL patients prior to treatment that in each case began to decline within a few days of the start of their anti-leishmanial drug therapy [128], and thus can be used as a marker of treatment response as well as a measure of parasitic burden over time. Recently, Hossain et al. [129] evaluated the use of real-time PCR and revealed the difference in parasite loads between primary VL and relapse VL. Subsequently, in a larger cohort of asymptomatic subjects, we established the threshold of parasitemia (> 5 L. donovani parasite genomes detected/mL) in blood for clinical symptoms of VL to occur [61]. Later, in an enlarged cohort of 1606 healthy individuals, of whom 442 were recent sero-converters with DAT and/or rK39, the risk for progression of disease was found to be much higher in qPCR-positive patients (odds ratio 14.8, 95% CI 5.1–42.5) (Chakravarty et al., personal communication).

Table 2 Summary of the comparative analytical sensitivity of real-time polymerase chain reaction assays targeting the Leishmania DNA region
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Elevated levels of interleukin (IL)-10 during active disease is a hallmark of VL, and this overproduction of IL-10 promote parasite replication and disease progression. Verma et al. [145] evaluated the parasitic burden measured by qPCR and its association with IL-10 production in VL and found that high qPCR load strongly correlates with plasma IL-10 levels, making it suitable for a biomarker of disease severity. Later, Wilson’s group developed several qPCR methods and strategies for Leishmania species differentiation and quantification in clinical specimens [63]. Leon et al. [146] evaluated the analytical performance of qPCR methods (designed on primers directed at kDNA, HSP70, 18S, and ITS-1 targets) and found that the 18S marker presented the highest sensitivity and specificity [146].

A qPCR assay usually provides a measure of the parasite load of blood at a given timepoint, but it remains unclear how this load can be correlated to the load at infection because the parasite load may vary with time, and likely reflects both host parasite interactions as well as the initial load. A number of researchers use PCR-ELISA for early detection and quantification, which allows multiple sample testing using whole blood, with a sensitivity of 87% [101, 147]. However, this method is tedious, expensive, and less sensitive than qPCR and has been tested on a limited number of clinical samples [101, 148].

2.3 Species Identification

VL is the most common Leishmania disease in the Indian subcontinent; however, recent identification of CL patients in Rajasthan (caused by L. tropica) and Himachal Pradesh (caused by L. donovani and L. tropica) [149,150,151] suggest that the clinical profile of CL is different in these states. Therefore, species identification assays are useful in such areas for proper management of the control programs. Importantly, in the Indian subcontinent (mainly India and Bangladesh) as well as in Africa (mainly Sudan), where L. donovani is the causative parasite for VL, a common complication of VL is PKDL [152]; it occurs in up to 50% of people who have recovered from VL in the months following treatment. It is much less common in India, with an incidence of less than 5–10%, and when it does occur, it does so many years after the acute infection [153]. In Africa, PKDL is even more common, but there are important intra-regional differences. It is most common in Sudan: up to 50–60% of VL cases develop PKDL, usually within 6 months, and virtually all cases develop within 12 months (mean 4.5 months). In Ethiopia, Kenya, and Uganda, PKDL is less common for reasons that are not well-understood.

Through whole-genome sequencing, Downing et al. [71] reported that there are a large number of chromosome copy number variations between L. donovani strains and other Leishmania species on the Indian subcontinent [71]. Therefore, better characterization of the parasite strain (i.e., species differentiation) is needed to resolve the mystery of whether the disease is due to reactivation of persistent parasites following clinical cure of VL or re-infection, and also to establish the cause of different forms of PKDL.

Commonly used target genes in Leishmania for species identification includes ITS (non-coding spacer DNA located between the 18S rRNA and 5.8S rRNA) [52, 154,155,156], repetitive nuclear DNA sequences [157], cytochrome-b genes [158, 159], mini-exon genes [160], G6PD genes [161], cpb genes [162, 163], gp63 genes [164], and hsp70 genes [165, 166]. For example, digestion of the ITS-1 PCR product with the Hae-III restriction enzyme differentiates most of the Leishmania species. The restriction fragment length polymorphism (RFLP) pattern is dependent on the restriction enzyme used, and thus the use of sequencing for confirmation is suggested. The random amplified polymorphic DNA technique (RAPD) is another molecular assay where amplification of DNA is performed using arbitrarily short primes without knowing the target sequences. Several studies have been performed using RAPD for investigation of genomic diversity [167,168,169,170], but its use in leishmaniasis is restricted due to the need for specific PCR standardization conditions and poor reproducibility [171]. Amplified fragment length polymorphism (AFLP) is a more advanced assay for investigation of variations in strains or closely related species [172]. It uses a restriction enzyme for genomic DNA digestion followed by selective PCR amplification of restriction fragments. Recently developed more sensitive PCR–fingerprinting techniques include multilocus sequence typing (MLST), which is based on the PCR amplification of multiple unlinked housekeeping genes followed by sequencing [68]. Moreover, a multilocus microsatellite typing (MLMT) approach has recently been developed by which East-African strains of L. donovani and Mediterranean strains of L. infantum could be resolved and assigned to genetically isolated populations [57, 74]. Srivastava et al. [70] explored the discriminatory power of different molecular assay and markers to detect genetic heterogeneity in clinical isolates of L. donovani from India [70]. Multilocus enzyme electrophoresis (MLEE) is another technique based on protein-based method which differentiates Leishmania parasites to species and subspecies levels using the electrophoretic mobility of enzymes [173]. This method has been known as the gold standard for characterization and identification of parasite strains. However, the requirement of mass cultivation of parasites, its low differentiation power in a homologous population, and the development of more sensitive molecular markers as alternative methods are the major drawbacks of MLEE [174]. Hernandez et al. [175] identified six new world Leishmania species through implementation of a High Resolution Melting (HRM) genotyping assay, which is another robust, highly sensitive, and reproducible genotyping technique.

2.4 Phylogenetic Analysis

The evolutionary pattern among species and the taxonomic status of Leishmania parasites are essential to understand the divergence among closely related species, design reliable diagnostic tools, and develop novel control methods. The malaria field is driving much of the relevant technology for this type of work. A major limiting factor to leishmaniasis is a critical lack of expertise throughout endemic areas. So far, many Leishmania strains have been typed by MLEE. On the other hand, introduction of numerous molecular typing methodologies with multicopy targets or multigene families have improved the analysis of phylogenetic, taxonomic, and genetic studies. These include DNA targets such as ITS [176], the single copy gene for the catalytic polypeptide from DNA polymerase α (polA) [177], cytochrome oxidase II (COII) gene [178, 179], cpB genes [180], 7SL RNA [181], and, most recently, the hsp70 subfamily sequence [165]. For example, Zhang et al. [73] investigated the phylogenetic relationship using ITS1 and kinetoplast COII gene sequencing and hypothesized that the phylogeny of Chinese Leishmania strains is associated with the geographical origin rather than the clinical form of the disease [73]. Fraga et al. [77] analyzed the phylogenetic study of 43 Leishmania strains from different geographic origins using the hsp70 sequence and found that monophylactic genus Leishmania consisted of three distinct subgenera: L. (Leishmania), L. (Viannia), and L. (Sauroleishmania) [77].

3 Technical Challenges and Future Prospects of Molecular-Based Assays

VL-affected patients living in endemic areas will not have access to quality care unless efforts are made to integrate existing innovative diagnostic technology into clinical management. For example, 3520 VL cases were reported in Sudan in 2014 and only 62% were diagnosed as confirmed VL. Molecular diagnostics are not only beneficial for patients, but if carried out through active case detection (such as sero surveys with the rk39 RDT) in the villages it will also reduce the parasite reservoir in highly endemic areas, given that humans are the only host reservoir for L. donovani on the Indian subcontinent. Though the sensitivity and diagnostic accuracy of molecular assays are reasonable to excellent in laboratory-based evaluations (reference laboratories), these methods are not currently able to be adapted to a PHC setting due to the expensive infrastructure and technical expertise required. The overall cost associated with a PCR assay is less than US$5 (230.0 Indian rupees) per sample [44]. Though this cost is two times greater than that for the rK39 RDT, it is currently not clear how such innovative techniques will replace rk39 RDT testing and how it can be meaningfully applied within the health system context of VL endemic areas. However, it is assumed that such rapid and highly sensitive molecular tools to assist clinicians working in the frontline PHC setting will help to better manage patients presenting with fever-related clinical syndromes. Strengthening early diagnosis and treatment capacities in PHC settings may provide long-term sustainability of the elimination effort through integrated case management as close as possible to the patient’s village. Importantly, the emergence and spread of drug resistance is a challenge for the VL control program. Therefore, monitoring drug efficacy and early reporting are essential as the drug arsenal is limited. Molecular detection tools would constitute a more rapid and high-throughput alternative to detect drug-resistant parasites, but requires a standardized way to use them and a structure to implement them in the sentinel sites.

4 Conclusion

There is no preventive or therapeutic vaccine for VL and the arsenal of anti-leishmanial drugs is limited; therefore, it is important to identify VL patients likely to relapse after drug treatment as well as new ways to recognize individuals who have had recent exposure to live parasites. Effective clinical management, chemotherapy, and control of transmission depend largely on early and unequivocal diagnosis. Molecular-based methods that can detect infection at a low level have recently become popular tools for diagnosis, and are relevant to the goal of VL elimination. To date, several molecular-based assays have been developed and evaluated, but PCR-based assays are found to be simple, rapid, and highly sensitive. The availability of such rapid tests that can be used to diagnose VL and as a marker of cure at peripheral health centers could have a great impact on the way VL is managed in endemic communities. These tests could be an alternative to the current rK39 dipstick test for accurate diagnosis and could be used to identify treatment failures and relapses.