Diagnostic advances in tegumentary leishmaniasis: a narrative review from 2018 to 2023

Introduction

Background

The urbanization process, the variability of climate change, trade, human movement, pollution, misuse of land, water storage, and irrigation, are all global aspects that can influence the incidence of vector-borne diseases. Although it is difficult to accurately predict how these changes will evolve, it is estimated that by 2030, 46 billion people will be living in urban areas around the world (1). This change will become a risk factor for the genesis of epidemic outbreaks of leishmaniasis, one of the 13 neglected tropical diseases (NTD) which are characterized by a potential loss of years of life also known as disability-adjusted life year (DALY) (2). This is especially true in endemic areas (3) of tropical and subtropical zones and is distributed in 98 countries in Europe, Africa, Asia, and America (4). Leishmaniasis is a parasitic disease caused by protozoa of the Trypanosomatidae family of the genus Leishmania, subgenus Leishmania and Viannia (5), and is transmitted to animals and humans by vectors of the Psychodidae family. In its entirety, the genus includes 31 species, 22 of which infect humans (5-7).

During its phase in the vertebrate host, these parasites can produce alterations in the skin, mucous membranes, and cartilage, giving place to the tegumentary form known as cutaneous leishmaniasis (CL), mucocutaneous leishmaniasis (MCL), diffuse cutaneous leishmaniasis (DCL), disseminated cutaneous leishmaniasis (DsCL), leishmaniasis recidivans (LR) and post-kala-azar dermal leishmaniasis (PKDL) (8). They can also affect organs such as the liver, spleen, and bone marrow, giving rise to the visceral form [visceral leishmaniasis (VL)] with its own particular set of symptoms and complications (3). In the Old World, tegumentary leishmaniasis is caused by five species of Leishmania: L. (L.) infantum, L. (L.) tropica, L. (L.) major, L. (L.) aethiopica, and L. (L.) donovani, while in the New World it is caused by the two subgenus Leishmania (L.) and Viannia (V.), such as Leishmania (L.) amazonensis, Leishmania (L.) mexicana, Leishmania (V.) braziliensis, and Leishmania (V.) guyanensis, that are endemic in Central and South America (5,9).

According to the World Health Organization (WHO), the prevention and control of leishmaniasis requires a combination of intervention strategies such as (I) early diagnosis and appropriate care; (II) vector control; (III) effective surveillance of the disease; (IV) control of animal reservoirs; and (V) social mobilization and strengthening of alliances (10). There exists a lack of adequate resources for leishmaniasis diagnosis especially in endemic areas, therefore the detection and correct identification of the parasite species remains a significant challenge. Current available diagnostic methods include parasitological, immunological, and molecular methods. The most accessible diagnostic methods, particularly in endemic areas, do not include a way to identify the species responsible for the disease. The diagnosis of VL, which is fatal if untreated, relies on a combination of clinical signs and parasitological, serological, and molecular tests (11). Serological tests have limited value when diagnosing cutaneous and MCL due to their limited availability in most countries, as well as the possible lack of sensitivity which is associated with a poor humoral response, therefore parasitological tests are used to confirm diagnoses (8). Parasitological tests, thanks to their great specificity, remain the reference standard (gold standard) in the diagnosis of CL although some studies showed that when these are associated with molecular tests the sensitivity and specificity are much higher (12). They are used to corroborate the clinical manifestations (13) and have become particularly useful in regions of the New World, where several species of Leishmania coexist with diverse clinical outcomes and responses to treatment (14). The Pan American Health Organization (PAHO) has developed in the Americas the Neglected, Tropical and Vector Borne Diseases Unit for the surveillance, strengthening, and application of leishmaniasis control measures. Despite its effort, 252,998 cases of tegumentary leishmaniasis were reported with an annual average of 42,166 during the implementation period of the Action Plan on Leishmaniasis between 2017 and 2022. Of these, 97% occurred in the Andes (41%), Brazil (37%) and Central America (19%) (15). Even when a 24% reduction in the number of cases in the region was observed, except in Guatemala, Mexico, and Panama, the disease was far from being controlled (15).

We carried out this narrative review due to the diagnostic limitations of tegumentary leishmaniasis to learn about new diagnostic methods for this NTD that have been recently published and that could possibly be candidates for approval of use and commercialization in endemic countries.

Rationale and knowledge gap

Unlike VL, which has serological and molecular methods that can be used as control tools for early diagnosis and timely treatment, tegumentary leishmaniasis still only has parasitological and molecular methods as the gold standard for diagnosing. Of these, the former have a low sensitivity due to a lack of training, poor sample quality due to small amounts of parasite being collected, inadequate implementation of the technique used to maintain the parasites in vitro, contamination, and inadequate infrastructure. In addition, there are limitations of microscopic diagnosis by smear, due to its low sensitivity and the inability to recognize biological species. The species are very diverse and, in some regions, it is necessary to identify the specific species to be able to administer the appropriate treatment.

Although molecular diagnostic methods have better sensitivity and specificity, their development presents limitations such as the implementation of the necessary infrastructure and the proper training of personnel at the field level. Immunological diagnostic methods show little development for tegumentary leishmaniasis.

Objective

The aim of this review is to present the current state of new technologies related to human tegumentary leishmaniasis diagnosis, in order to identify future innovations that may be candidates for improving early diagnosis of tegumentary leishmaniasis. We present this article in accordance with the Narrative Review reporting checklist (available at https://jphe.amegroups.com/article/view/10.21037/jphe-24-51/rc).

Methods

Using the PubMed database from 2018 to June 2023, peer-reviewed articles in English were selected. The search was driven by title and abstract for further evaluation. The terms used to search for articles related to tegumentary leishmaniasis diagnosis were: (“Leishmania”) AND (“diagnosis*”) AND (“specificity” OR “sensitivity”) NOT (“visceral leishmaniasis”) NOT (“dog”) (Table 1). The articles inclusion criteria were: the publication has been peer-reviewed and published in English. Exclusion criteria were: (I) review articles; (II) VL diagnostic studies; (III) PKDL; (IV) T. cruzi; (V) animal studies.

Data extraction was performed by 10 authors and one author checked its accuracy. For study characteristics, we extracted: title, abstract, authors’ names, year, journal name, number, volume, pages, sensitivity, specificity, Leishmania parasite species, clinical diagnosis, country and DOI and then applied the exclusion criteria.

Results

A narrative review was performed. Of the 87 articles found, 33 were deemed relevant and included in the study and divided in four tables: parasitological diagnostics methods (Table 2); immunological diagnostic methods (Table 3); molecular diagnostic methods (Table 4) and new approaches in diagnostics methods for tegumentary leishmaniasis (Table 5).

Discussion

CL is considered one of the most neglected diseases of great importance as shown by Bharadwaj et al. (2), which is associated with poverty. Review results will be discussed as follows.

Parasitological diagnostic methods for tegumentary leishmaniasis

Direct parasitological diagnosis methods for tegumentary leishmaniasis are shown in Table 2. Studies proved that the gold standard parasitological diagnostic method, smear (S), achieves a sensitivity between 40% to 78.97% for the diagnosis of CL when compared with the molecular methods recombinase polymerase amplification (RPA), conventional polymerase chain reactions (cPCR) amplifying molecular targets such as kinetoplast DNA (kDNA) or heat shock protein 70 (HSP70). This is in countries such as Colombia (16), Brazil (17) and Afghanistan (18), with a specificity of 77.27% in the latter (18) (Table 2). In tegumentary leishmaniasis diagnosis, parasitological techniques remain the gold standard (48), which, although are highly specific for diagnosing, are insufficiently sensitive (49) as also observed in this review (16-18).

We also found that Ferreira et al. (19) in Brazil, used the histopathological method (HP) to observe the parasite, showing a sensitivity of 50% when compared to the traditional culture method (TCM) in samples of CL by L. (V.) braziliensis (Table 2). The histopathological technique as a diagnostic method is used but has a low sensitivity (50), as observed by Ferreira et al. (19).

With regards to the TCM, another gold standard method for CL diagnosis, two studies in French Guiana (20) showed a sensitivity between 49% and 76% in CL by L. (V.) braziliensis, while Tawfeeq et al. (21) showed that TCM reaches a sensitivity and specificity of 80.77% and 100%, respectively, in identifying L. (L.) major, when compared to cPCR and nested-PCR (nPCR). In contrast, the microculture method (MCM) (Table 2) showed a sensitivity and specificity of 95% and 97.8%, respectively, when compared to TCM in the diagnosis of CL by L. (L.) aethiopica in Ethiopia (22). Concerning the sensitivity of the traditional culture technique, they have variable ranges as well as S (20,21). This variability is partly explained by the limitations of the technique, such as the need for specialized infrastructure to maintain axenic cultures, the high cost, the time required, the specialized personnel and others (50). However, the combination of S and culture increases the diagnostic sensitivity for CL as well as having a high specificity (100%) (21). Its advantages lie in the increased sensitivity and the fact that it allows isolation of parasite strains, which can be used for multiple subsequent analyses. Similarly, high sensitivity and specificity were observed by Aberra et al. (22) using the microcapillary culture method, as well as by Pagheh et al. in Iran (51). Nevertheless, the same technical limitations exist for MCT.

While with proper infrastructure and adequate technical training, gold standard methods are not the ideal methods to use in endemic countries where leishmaniasis is a NTD, and instead immunological and molecular assays have to be used to confirm the diagnosis.

Immunological diagnostic methods for tegumentary leishmaniasis

Immunohistochemistry (IHC) has emerged as an affordable alternative to PCR for LC diagnosis. Some laboratories recently used monoclonal antibodies anti-CD1a against Leishmania (52), the goal was to test the anti CD1a antibody in at least 1 case with a very low parasitic index, those were promising results in the application of this type of monoclonal antibody in the IHC technique. There are two hypotheses about the use of monoclonal antibody against CD1a despite is a mammalian protein. The first one is that, during exocytosis, amastigotes forms acquire CD1a; and the second one is by cross-reactivity, which is the least accepted. In studies of Immunological based diagnostic methods for tegumentary leishmaniasis (Table 3), such as IHC done in Brazil, Ferreira et al. (19) showed 66% of sensibility when compared with TCM in CL samples caused by L. (V.) braziliensis, whereas Sánchez-Romero et al. (23) reported an 82.3% sensitivity when compared to cPCR for MCL by L. (V.) braziliensis and L. (L.) amazonensis diagnosis. On the other hand, Freire et al. (24) observed samples of CL by L. (V.) braziliensis, L. (L.) amazonensis and L. (L.) guyanensis, and got an 87.5% and 90% sensitivity and specificity respectively when compared with cPCR, Montenegro test, Biotin hyperimmune sera by enzyme-linked immunosorbent assay (ELISA) and quantitative polymerase chain reaction (qPCR). In Brazil (19,23,24), the studies showed a great sensitivity (Table 3) but mentioned limitations were caused by the lack of method standardization, in addition to the sample processing risk of damaging the tissue and antigens which could affect the sensitivity.

A study conducted in Tunisia by Saide et al. (25) used direct immunofluorescence (DIF) (Table 3) and detected infected mononuclear phagocytic cells and Leishmania amastigotes using FITC-labeled anti-L. (L.) major IgG with 98.3% sensitivity and 100% specificity when compared with the gold standard (S) and kDNA qPCR. Conversely, Stensvold et al. (26) (Table 3) compared the sensitivity and specificity of commercial indirect fluorescent antibody technique (IFAT) and ELISA for CL by L. (V.) braziliensis, L. (V.) guyanensis, L. (V.) panamensis, L. (L.) major, L. (L.) mexicana and L. (L.) tropica diagnosis. Both methods presented 95.5% sensitivity for CL diagnosis, but IFAT offered 42.9% specificity when compared with ELISA and Western blot, whereas ELISA’s sensitivity was 81% when compared with IFAT. Regarding the DIF and IFAT, Saidi et al. (25) showed that DIF has higher sensitivity and specificity (Table 3). In the previous study (53), this same group already reported that this method could detect low parasite load that was not detected by microscopy but was by conventional internal transcribed spacer 1 (ITS1) PCR, becoming a new candidate for second line diagnostic confirmation. Concerning IFAT, Stensvold et al. (26) also showed in their study that it has a great sensitivity although the specificity was low, the latter being one of the disadvantages of indirect IFA since it presented cross-reactivity with other diseases caused by trypanosomatids such as Chagas disease (53). As such, it is not recommended in countries where both diseases coexist.

Regarding ELISA (Table 3), in the new world Bracamonte et al. (27) used membrane crude antigens (MCAs) and reported 98% sensitivity and 63.6% specificity in CL by L. (V.) braziliensis diagnosis when compared to the gold standard (S), Leishmania skin test (LST) and a molecular method known as polymorphic specific polymerase chain reaction (PS-PCR LST). In Sri Lanka, De Silva et al. (28) (Table 3) used In-House ELISA for CL by L. (L.) donovani diagnosis. When comparing it with nPCR, which amplifies the ITS1, they observed 84.4% sensitivity and 50% specificity. Recently, Viana et al. (29) in Brazil, using a chemiluminescent ELISA (cELISA) (Table 3) reached 94% sensitivity and 97% specificity when compared with conventional PCR and ELISA for CL by L. (L.) braziliensis and L. (L.) major diagnosis. In the assays using ELISA for CL diagnosis, the sensitivity and specificity depends on the antigens used (54). Duarte et al. (55) evaluated serum from patients with CL and ML by ELISA using recombinant proteins such as tryparedoxin peroxidase from L. (V.) braziliensis, which showed 100% sensitivity and specificity. On the other hand, serological studies using crude antigens from amastigotes and promastigotes membranes (MCA) like those done by Bracamonte et al. in 99 subjects diagnosed as ATL, 27 as no ATL, and 84 donors from non-ATL-endemic areas (27) showed (Table 3) high sensitivity but low specificity, increasing to 98.4% (94.4–100%) when sera with anti-Trypanosoma cruzi antibodies that displayed cross-reactivity were excluded. This turns it into a good candidate for tegumentary leishmaniasis diagnosis in seroprevalence analysis. There are some evidence in the literature about the peroxidoxin in L. (V.) braziliensis as a candidate for a diagnosis for TL. The data suggested that peroxidoxin has a promising potential to identify TL, VL and CVL (56). However, are necessary prospective studies from endemic areas to better characterize this approach. In Sri Lanka, De Silva et al. (28) assessed a new inhouse ELISA demonstrating a high sensitivity but a low specificity when compared with ITS1 nested PCR, increasing its sensitivity to 99% when combining slit-skin-smear (SSS) and serum ELISA, and thus favoring it as a complementary diagnostic test. Stensvold et al. (26) using a commercial ELISA, NovaLisa™ Leishmania infantum IgG, with a modified cut-off for borderline positive values, portrayed a high sensitivity and specificity in countries such as Denmark where, until a few years ago, they did not present any cases of CL. Similarly, the study by Viana et al. (29) showed high sensitivity and specificity with chemiluminescent ELISA using many glycoconjugates, which present antibodies with significantly high levels of anti-α-Gal IgG, NGP28b standing out among them, which suggests that anti-NGP28b antibodies could be candidates for early cure biomarkers for LC. In addition, the glycoconjugates expressed a low cross-reactivity with control patients’ sera that were infected with Chagas disease (53).

There are also studies that used rapid diagnostic tests (RDTs) for CL diagnosis (Table 3). Shalling et al. (30) using the Detect™ Rapid Test (CL Detect) displayed 35.8% and 36.7% sensitivity and 83.3% and 85.7% specificity for L. (L.) guyanensi in Surinami when compared with gold standard (S) and cPCR, whereas Aerts et al. (18), using the same kit in a cost-benefit study, showed 66.27% sensitivity and 95.45% specificity when compared with cPCR for CL by L. (L.) tropica diagnosis in Afghanistan. Similarly in Morocco, Bennis et al. (31) observed 68% sensitivity and 94% specificity when compared with gold standard (S), PCR-ITS and kDNA PCR for CL by L. (L.) tropica and L. (L.) major. In general, RDTs are quick and easy to perform and can be used in settings with limited laboratory infrastructure and staff (57). Several years after the successful development of an RDT for VL (58,59) an RDT for CL has been developed. The Low sensitivity observed in rapid assays with CL RDT (18,30,31) (Table 3), indicates that they could be used in primary screening, with the disadvantage of presenting high percentages of false negatives, needing confirmatory testing, and contrasting the findings using rapid immunochromatographic tests developed for VL with rK39 RDT, which offers 97% sensitivity in India and 85% sensitivity in Eastern Africa (60).

Molecular diagnostic methods for tegumentary leishmaniasis

The molecular diagnostic methods for tegumentary leishmaniasis (Table 4), such as cPCR, showed variable sensitivity ranging between 78% (16), 80% (17) and 100% (17) when targeting the molecular amplification of Leishmania (Viannia) DNA, HSP70, and kDNA from Leishmania spp. in patients with CL in countries such as Colombia, Brazil and Iran respectively. The molecular diagnostic methods also showed 92.8% (21) sensitivity when amplifying Leish1 and Leish2 antigens from L. (L.) braziliensis and L. (L.) amazonensis in MCL samples in Brazil, an 81% specificity for Leishmania spp. causing CL (16), and a 100% specificity for zoonotic cutaneous leishmaniasis (ZCL) by L. (L.) major in Iran (32).

Molecular methods have become more attractive as they offer sensitive, specific, reliable and rapid parasite detection, but their implementation requires specific material and well-equipped laboratories. Several molecular diagnostic tests for CL diagnosis have been developed and apparently, they portray higher sensitivity and specificity than traditional methods (61). The molecular targets for these methods are diverse (Table 4). For example, ones that are widely used are the amplification of kDNA from Leishmania spp. (17) and species-specific kDNA (SS-kDNA) from L. (L.) major in ZCL samples (32) or from L. (L). braziliensis and L. (L.) amazonensis pathogenic agents for MCL (23), all of which demonstrated high sensitivity. Cantanhêde et al. (17) showed that cPCR kDNA amplification presents greater sensitivity than HSP70 amplification as noted previously by other studies (62).

A high sensitivity was also observed amplifying kDNA using semi nested-PCR (34), possibly due to the repetitions presented in the amplified segment (63). In spite of having high sensitivity and specificity, molecular methods are expensive and are inaccessible in endemic areas.

PCR combined with high resolution melting (HRM) analysis (Table 4) which uses ITS as a molecular target showed 89% sensitivity and reached 100% specificity for L. (L.) major and L. (L.) tropica diagnosis in Iran (33). Similarly, in Brazil (34), HSP70 amplification using HRM demonstrated 96.4% sensitivity for CL and MCL diagnosis by L. (L.) amazonensis, L. (V.) braziliensis and L. (L.) guyanensis. In spite of showing high sensitivity and specificity, ITS and HSP70 amplification using qPCR-HRM (33,34) has the disadvantage of needing equipment and experienced personnel to perform Real Time PCR and HRM analysis, which are not accessible to all healthcare systems. In addition, the technique is susceptible to contamination, which can result in false positives if the necessary precautions are not taken during sample manipulation and analysis. On the contrary, for Asfaram et al. (33) this method decreases false positive results since it quantifies low-density nucleic acids, has a low probability of impurities, and, in epidemics, has the ability to diagnose quickly (90 min). Other molecular methods such as Duplex PCR for ITS1 amplification for American cutaneous leishmaniasis (ACL) diagnosis in Sri Lanka (35) portrayed a sensitivity and specificity of 83.06% and 86.96% respectively when it identifies Leishmania spp. species from Viannia subgenus and L. (L.) amazonensis. Cupolillo et al. (64) evaluated ITS from the Leishmania and Viannia subgenus from different hosts and geographic areas and demonstrated that ITS from this subgenus vary but that its rapid evolution allows us to distinguish between species.

On the other hand, Nateghi et al. in Iran (36) (Table 4) while amplifying ITS2 for CL by L. (L.) major, L. (L.) tropica and L. (L.) infantum diagnosis, observed 100% sensitivity and specificity. Similarly, in Iran, Khoshnood et al. (37) used the semi-nPCR to amplify the kDNA and observed 93% sensitivity and 100% specificity. The usage of loop mediated isothermal amplification (LAMP), amplifying 18SrRNA, for CL diagnosis by Schallig et al. (30) (Table 4) in Afghanistan, showed sensitivity of 84.8% and 91.4% and specificity of 42.9% and 91.7% for L. (L.) major, L. (L.) tropica, L. (V.) braziliensis, L. (L.) panamensis, L. (L.) mexicana and L. (L.) guyanensis diagnosis when compared with microscopy and GIEMSA stain (GS). Similarly, Aerts et al. (18) with the same molecular target and the same pathogens observed 89% sensitivity and 64.64% specificity. In Tunisia, by amplifying the cysteine protease B (cpb) gene from L. (L.) major and L. (L.) tropica, Chaouch et al. (38) found 84% and 100% sensitivity and specificity respectively, when compared to qPCR, conventional microscopy (CM), and conventional cpbPCR. Isothermal amplification methods have been developed in recent years using various molecular targets, such as 18srRNA (18,30) and cpg gene (38), with a high diagnostic sensitivity as previously observed by other authors (65). The technique should be monitored to avoid contamination issues with undesired amplifications due to its high sensitivity. One of its advantages is that it can process several samples at the same time, decreasing costs when used at its maximum capacity (18).

New approaches in diagnostic methods for tegumentary leishmaniasis

Table 5 shows a summary of the articles that demonstrate advantages for their applicability both in reference centers and in the field, the latter being the main interest because it is where diagnostic methods should be implemented for epidemiological purposes as stated by other authors (2).

Technological innovations, although far from being immediately applicable, show new diagnostic possibilities for the future. Regarding the ELISA’s use for diagnosis of CL and MCL by L. (V.) braziliensis in Brazil, García et al. (39) (Table 5) describe the advantages of the proteomic development of a chimeric protein identified as rCHIP, showing 100% sensitivity and 100% specificity when compared to both ELISA with the soluble antigen (SLB) from L. (V.) braziliensis and the rapid IFA. In the era of proteomics, it is worth highlighting the development of the chimeric protein rCHIP from the combination of 13 linear B cell epitopes, which was used in ELISA, presenting 100% sensitivity and specificity for both CL and MCL (39) (Table 5). The chimeric protein was able to identify antibodies with low titration, thus avoiding cross-reactivity which is observed in other ELISAs used for CL diagnosis (66). This type of study leads the way for technological innovations such as a new immunochromatographic assay that could help early diagnosis in countries with a lack of infrastructure.

Innovations in molecular techniques are described in two articles and involve assays performed with the isothermal recombinase RPA technique directed to the CL diagnosis in Colombia (16) and Ghana (67). Mesa et al. (16) showed 90.4% sensitivity and 72.7% specificity when compared to the gold standard (S) as well as a 72% sensitivity and 69.8% specificity when compared with cPCR. Yeboah et al. (40) used RPA together with lateral flow (RPA-LF) which showed a sensitivity of 88.9% when compared to qPCR for the confirmation of the presence of L. (L.) donovani and L. (L.) major in CL lesions.

In Colombia, Cossio et al. (67) combined isothermal amplification of Leishmania kDNA together with a lateral flow immunochromatographic strip, both in the field and in a reference center. They obtained samples with minimally invasive swabs and fast technology for analysis of nucleic acids (FTA) filter paper of lesions of over 2 weeks evolution. In the reference center, it showed a sensitivity and specificity of 87% [95% confidence interval (CI): 74–94%] and 86% (95% CI: 74–97%), respectively. In the field, sensitivity was 75% (95% CI: 65–84%) and specificity 89% (95% CI: 78–99%). Positive likelihood ratios in both scenarios were greater than 6, while negative likelihood ratios ranged from 0.2 to 0.3, supporting the utility of RPA-LF to potentially rule out infection. All of this, in addition to the low complexity of combined RPA-LF and non-invasive sampling, makes it a strong candidate for immediate application.

kDNA amplification through modified and optimized qPCR combined with immunomagnetic separation (IMS) assay for Leishmania promastigote capture showed 93% of positivity at 72 h versus 50% within 2–4 weeks incubation (TCM) for CL by L. (L.) major diagnosis in Tunisia (41). The high positivity showed the ability of identifying promastigotes at a low parasite load in early cultures. Although, IMS in combination with IFA, showed a lower efficiency, between 25–42%. In contrast, a study suggested that IMS in combination with qPCR can be a fast and sensitive method for Leishmania promastigote detection like it has been demonstrated for visceral Leishmania diagnosis in previous studies (68).

Comparably, studies employing PCR nucleic acid amplification technologies in portable equipment have been published for CL and MCL diagnosis in Peru by Kariyawasam et al. (42) who amplified kDNA and patented it as PALM-PCR^®. This study showed 100% (95% CI: 83.2–100%) sensitivity and 25% (95% CI: 3.2–65.1%) specificity in the field. The sensitivity of this method was calculated in an on-field study in which 20 samples were analyzed and 19 of them were confirmed using qPCR. Those results show that 19 of 20 samples (95%) of the population will get confidence interval in the same parameter. However, it would be more specific to provide the confidence interval, as the sample size was small (42). The advantage of portable PCR is the speed at which a diagnosis can be made, usually within an hour. The results in the field portrayed a low specificity, probably due to heat and humidity since the studies were conducted in the Peruvian amazon, resulting in false positives when compared with qPCR. It is an innovative method but its specificity has to be improved by correcting this technical issue. It is worth taking into consideration that it showed 90% (95% CI: 55.5–99.8%) sensitivity and 91.7% (95% CI: 77.5–98.3%) specificity in house. On the other hand, recently in Colombia and in Peru, Castellanos-Gonzalez et al. (43) used Mini PCR to conduct a study for CL by L. (V.) panamensis diagnosis which showed 100% specificity and sensitivity. SYBR gene amplification utilizing qPCR was preceded by a simple DNA extraction, using a buffer prepared in house, from L. (V.) panamensis promastigotes originating from Peruvian and Colombian patient samples with suspected CL obtained using filter paper. Field studies need to be carried out because the miniPCR^® test has a great potential to contribute to diagnosis in remote endemic areas, but it is necessary to test its sensitivity and specificity in the field.

Diagnostic studies that incorporated sanger sequencing (SS) of rRNA intergenic spaces (IGS) from MCL samples by L. (V.) brazilensis in Brazil were also found (Table 5) with a reaction efficiency of 98.08% (69). The SS, first described in 1977 (69), is considered a molecular technique with a high sensitivity and specificity for Leishmania spp. detection (36). Recently, Osorio-Peralta et al. in Colombia (70), using products from ITS-1 PCR amplification and sequenced by the Sanger method, showed a reaction efficiency of 99.6% with the homology between L. (L.) infantum and L. (L.) donovani species.

Additionally, Blaizot et al. (20) in French Guiana (Table 5), used the technique for sample extraction from CL patient’s ulcers with cotton swabs (CS), followed by qPCR with SYBR green and finishing with a SS using HSP70 from L. (V.) braziliensis as a molecular target. The results showed 98% sensitivity versus the gold standard (S), biopsy (B), TCM and PCR-restriction fragment length polymorphism (PCR-RFLP). All real-time PCR positive samples were successfully identified on a species level through DNA sequencing. Just like in Castellanos-Gonzalez et al. (43) and Blaizot et al. (20) in French Guiana, sample extraction with a CS is an excellent noninvasive extraction method that was also successfully used previously (71) for CL diagnosis through qPCR. A new approach for tegumentary Leishmania diagnosis involves using the CS noninvasive sample extraction technique, followed by HSP70 gene amplification through qPCR, which has already demonstrated to be highly sensitive for CL and MCL molecular diagnosis (36), and finally ending with sequencing using the Sanger technique, which has a high percentage of reaction efficiency. This also demonstrated a 100% homology of these genes within etiologic CL species like L. (V.) braziliensis (44).

Despite highly precise sequencing data enabled by first, second, and third generation sequencing technologies, short-read technologies remain limited due to PCR amplification and short read lengths. This technological limitation hinders progress for their use in clinical diagnostics. Third-generation technologies allow much longer read lengths, enabling more uniform and direct sequence evaluation while circumventing the limitations of short-read technologies. The primary limiting factor of these third-generation technologies was a high error rate, making them clinically inadequate. The ongoing refinement of these third-generation technologies and bioinformatics tools to enhance accuracy are promising for the next wave of impactful advances. However, expecting short and long-term application of these technologies in endemic countries for LC is a utopia, given the cost of equipment and the need for trained human resources in bioinformatics analysis (72).

Chakkumpulakkal et al. (45) described a new approach for CL diagnosis using synchrotron-based Fourier-transform infrared (SR-FTIR) microspectroscopy, a bioanalytical and imaging tool first described by Holman et al. (73). Although the current study employs an expensive synchrotron-based (FTIR) microspectroscopic approach, it serves as a proof-of-concept for Leishmania diagnosis using infrared spectroscopy on aqueous samples, allowing researchers to locate, identify, and track specific chemical events within individual live mammalian cells. Mid-IR photons have too low energy (0.05–0.5 eV) to break bonds or cause ionization, indicating that the synchrotron IR beam has no detectable effects on short- or long-term viability, reproductive integrity, cell cycle progression, or mitochondrial metabolism in live human cells, and only minimally heats the sample (<0.5 ℃). These studies have laid a significant foundation for SR-FTIR in biological and biomedical research. In the case of L. (L.) major infection, SR-FTIR was used to analyze the infrared radiation emission from lipids present in the membranes of 160 functional infected aqueous and 130 uninfected macrophages, as well as 35 amastigotes and 80 promastigotes (45). Lipidomic profiling of L. (L.) major showed relative intensities of lipid and fatty acid bands in infected cells, potentially explained by higher amounts of glycolipids in the parasite’s membranes and organelles, which may affect parasite response to treatment and its ability to infect hosts (74). Analysis of results utilized algorithms for multivariate analysis of reflectance spectra including partial least squares-discriminant analysis (PLS-DA), support vector machine-discriminant analysis (SVM-DA), and k-nearest neighbors (KNN). Multivariate analyses of synchrotron radiation from infected and non-infected macrophages in physiological solutions showed sensitivities of 0.923, 0.981, and 0.989, respectively, and specificities of 0.897, 1.00, and 0.975, respectively. PLS-DA models with cross-validation in living amastigotes and promastigotes showed 98% sensitivity and specificity in the lipid region, and 100% sensitivity and specificity in the fingerprint region (1,800–1,000 cm⁻¹), indicative of contributions from proteins and nucleic acids. This fingerprint region in an infrared spectrum is unique for each compound (73). The study demonstrated the diagnostic potential of FTIR spectroscopy by identifying unique diagnostic bands that could potentially predict Leishmania infection in the future. The FTIR technique using synchrotron infrared microscopy, demonstrating that chemical compounds within cells can be visualized via diffraction. However, it’s application warrants future validation in both field and laboratory settings.

Welearegay et al. (46) published an innovative diagnostic method involving the analysis of volatile organic compounds (VOCs) in exhaled breath. They utilized a set of custom-designed chemical gas sensors based on ligand-capped CuNPs, demonstrating high sensitivity in detecting volatile gases in CL carriers in Tunisia. The study involved 28 volunteers diagnosed with CL and a control group of 32 individuals recruited from various endemic sites. The sensor test achieved a classification success rate for human CL with 98.2% accuracy, 96.4% sensitivity, and 100% specificity. Notably, one sensor, utilizing CuNPs functionalized with 2-mercaptobenzoxazole, showed perfect discrimination with 100% accuracy, sensitivity, and specificity for human CL. This technique holds promise for various health applications, as previously noted (75). Although the molecular basis of this method remains unclear, it potentially involves the peroxidation of polyunsaturated fatty acids, though the exact mechanism has yet to be elucidated.

Genomic editing through CRISP-Cas12 technology, taking DNAr18S and HSP70 as molecular targets for CL and MCL by L. (V.) braziliensis diagnosis, were carried out in Perú and other Latin American countries (47) (Table 5), establishing an innovative diagnostic tool for nucleic acid detection with high sensitivity and specificity. This tool could possibly be used out in the field in endemic areas in the future. The 18S PCR/CRISPR assay detected all the species without discrimination and without cross-reactivity with T. cruzi, unlike kDNA PCR/CRISPR which was only specific for L. (Viannia). This technique was performed with viral pathogens (76,77). The validation of this method should be performed in field assays, taking advantage of its high precision to detect Leishmania species even in the presence of a single parasite during each reaction and in recent lesions that have less than 3 months of evolution. This turns it into an excellent diagnostic alternative that can be used in the point of care (POC), which is also stated by Bharadwaj et al. (2).

Conclusions

Of the 87 articles found, 33 were deemed relevant and included in the study. These were divided in four groups: direct based parasitological, immunological based, molecular diagnostic based and new approaches. Data collected were about sensitivity, specificity, clinical manifestation, causative species, and country. Regarding sensitivity, molecular techniques like cPCR, qPCR, LAMP, duplex PCR, PALM PCR, HRM PCR, Mini-PCR, PCR-FT, Semi-NESTED PCR, and RPA-LF showed higher sensitivity immunological methods, such as ELISA and IHC, which in turn has higher sensitivity than direct parasitological methods. Serological tests could be an alternative for the diagnosis of CL by using the appropriate antigen and method, overcoming the limitations of sensitivity associated with a poor humoral response. Hence, a diagnosis is confirmed with Parasitological tests, which are still the reference standard in the diagnosis of CL, and when associated with molecular tests show higher sensitivity and specificity. Both culture and molecular diagnosis require considerable infrastructure and technical expertise, restricting their use to reference laboratories, making it difficult for a timely diagnosis to be made and thus for treatment to be initiated. As diagnostics advance, the chemiluminescence assay is a technique worth considering for the diagnostic confirmation of tegumentary leishmaniasis due to its high specificity and sensitivity (≥1 pg/mL), ease of automation, and potential for implementation in countries where ELISA is already established. In contrast, rapid immunochromatographic assays have low sensitivity and specificity, which is why, unlike with VL, its application for epidemiological trials is still distant. The isothermal RPA showed high sensibility. Moreover, the Palm and Mini PCR, achieved the highest sensitivity, showing themselves as strong candidates for a points of care epidemiological diagnosis, which is where these patients with suspected CL should be identified. Additionally, combining the recollection of samples using CS with qPCR and sequencing improves these techniques’ sensitivities when associated with the traditional methods used in leishmaniasis endemic countries. Finally, the CRISPR-Cas12a technique opens up a potentially new diagnostic alternative. As mentioned by Bharadwaj et al. (2), to achieve epidemiological control of neglected diseases, diagnoses must be made at the points of care of endemic areas, where there are major sanitary necessities but limited resources.

This revision puts forward different candidates that could serve as first line diagnostic techniques for tegumentary leishmaniasis, not just at a point of care level as stated by Bharadwaj et al. (2), but also in the reference centers. It is also worth emphasizing that the eradication of leishmaniasis and other neglected diseases must be approached from an interdisciplinary and multisectorial angle that should include strategic planning, multilevel integration of existing interventions, and the involvement of stakeholders at both policy making and community levels (2).

Acknowledgments

To Santiago Gangotena Gonzalez (In memory) for guiding us through liberal arts.

Funding: This work was supported by Escuela de Medicina, USFQ, Grant 2022 (No. HUBI17715).

Footnote

Provenance and Peer Review: This article was commissioned by the Guest Editor (Mellissa Withers) for the series “Equity in Health: Findings from the APRU Global Health Conference 2023” published in Journal of Public Health and Emergency. The article has undergone external peer review.

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Conflicts of Interest: All authors have completed the ICMJE uniform disclosure form (available at https://jphe.amegroups.com/article/view/10.21037/jphe-24-51/coif). The series “Equity in Health: Findings from the APRU Global Health Conference 2023” was commissioned by the editorial office without any funding or sponsorship. The authors have no other conflicts of interest to declare.

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References

1. United Nations Population Division. World urbanization prospects: the 2001 revision. Geneva: United Nations; 2002.
2. Bharadwaj M, Bengtson M, Golverdingen M, et al. Diagnosing point-of-care diagnostics for neglected tropical diseases. PLoS Negl Trop Dis 2021;15:e0009405. [Crossref] [PubMed]
3. Sutherst RW. Global change and human vulnerability to vector-borne diseases. Clin Microbiol Rev 2004;17:136-73. [Crossref] [PubMed]
4. Alvar J, Vélez ID, Bern C, et al. Leishmaniasis worldwide and global estimates of its incidence. PLoS One 2012;7:e35671. [Crossref] [PubMed]
5. Comité de Expertos de la OMS sobre el Control de las Leishmaniasis & Organización Mundial de la Salud. (2012). Control de las leishmaniasis: informe de una reunión del Comité de Expertos de la OMS sobre el Control de las Leishmaniasis, Ginebra, 22 a 26 de marzo de 2010. Organización Mundial de la Salud. Available online: https://iris.who.int/handle/10665/82766
6. Shaw JJ. Taxonomy of the genus Leishmania: present and future trends and their implications. Mem Inst Oswaldo Cruz 1994;89:471-8. [Crossref] [PubMed]
7. Ashford RW. The leishmaniases as emerging and reemerging zoonoses. Int J Parasitol 2000;30:1269-81. [Crossref] [PubMed]
8. de Vries HJ, Reedijk SH, Schallig HD. Cutaneous leishmaniasis: recent developments in diagnosis and management. Am J Clin Dermatol 2015;16:99-109. [Crossref] [PubMed]
9. Burza S, Croft SL, Boelaert M. Leishmaniasis. Lancet 2018;392:951-70. [Crossref] [PubMed]
10. Leishmaniasis. Available online: https://www.who.int/es/news-room/fact-sheets/detail/leishmaniasis
11. Sundar S, Rai M. Laboratory diagnosis of visceral leishmaniasis. Clin Diagn Lab Immunol 2002;9:951-8. [PubMed]
12. Bezemer JM, Merckx J, Freire Paspuel BP, et al. Diagnostic accuracy of qPCR and microscopy for cutaneous leishmaniasis in rural Ecuador: A Bayesian latent class analysis. PLoS Negl Trop Dis 2023;17:e0011745. [Crossref] [PubMed]
13. Reimão JQ, Coser EM, Lee MR, et al. Laboratory Diagnosis of Cutaneous and Visceral Leishmaniasis: Current and Future Methods. Microorganisms 2020;8:1632. [Crossref] [PubMed]
14. Carstens-Kass J, Paulini K, Lypaczewski P, et al. A review of the leishmanin skin test: A neglected test for a neglected disease. PLoS Negl Trop Dis 2021;15:e0009531. [Crossref] [PubMed]
15. Leishmaniasis: Epidemiological Report for the Americas 2023;12:14. Available online: https://iris.paho.org/handle/10665.2/59155
16. Mesa LE, Manrique R, Robledo SM, et al. The performance of the recombinase polymerase amplification test for detecting Leishmania deoxyribonucleic acid from skin lesions of patients with clinical or epidemiological suspicion of cutaneous leishmaniasis. Trans R Soc Trop Med Hyg 2021;115:1427-33. [Crossref] [PubMed]
17. Cantanhêde LM, Mattos CB, Cruz AK, et al. Overcoming the Negligence in Laboratory Diagnosis of Mucosal Leishmaniasis. Pathogens 2021;10:1116. [Crossref] [PubMed]
18. Aerts C, Vink M, Pashtoon SJ, et al. Cost Effectiveness of New Diagnostic Tools for Cutaneous Leishmaniasis in Afghanistan. Appl Health Econ Health Policy 2019;17:213-30. [Crossref] [PubMed]
19. Ferreira LC, Quintella LP, Schubach AO, et al. Comparison between Colorimetric In Situ Hybridization, Histopathology, and Immunohistochemistry for the Diagnosis of New World Cutaneous Leishmaniasis in Human Skin Samples. Trop Med Infect Dis 2022;7:344. [Crossref] [PubMed]
20. Blaizot R, Simon S, Ginouves M, et al. Validation of Swab Sampling and SYBR Green-Based Real-Time PCR for the Diagnosis of Cutaneous Leishmaniasis in French Guiana. J Clin Microbiol 2021;59:e02218-20. [Crossref] [PubMed]
21. Tawfeeq HM, Ali SA. Highly sensitive nested polymerase chain reaction to improve the detection of Leishmania species in clinical specimens. J Parasit Dis 2022;46:754-63. [Crossref] [PubMed]
22. Aberra L, Abera A, Belay T, et al. Evaluation of microcapillary culture method for the isolation of Leishmania aethiopica parasites from patients with cutaneous lesions in Ethiopia. Diagn Progn Res 2019;3:4. [Crossref] [PubMed]
23. Sánchez-Romero C, Júnior HM, Matta VLRD, et al. Immunohistochemical and Molecular Diagnosis of Mucocutaneous and Mucosal Leishmaniasis. Int J Surg Pathol 2020;28:138-45. [Crossref] [PubMed]
24. Freire ML, Rego FD, Lopes KF, et al. Anti-mitochondrial Tryparedoxin Peroxidase Monoclonal Antibody-Based Immunohistochemistry for Diagnosis of Cutaneous Leishmaniasis. Front Microbiol 2021;12:790906. [Crossref] [PubMed]
25. Saïdi N, Galaï Y, Ben-Abid M, et al. Imaging Leishmania major Antigens in Experimentally Infected Macrophages and Dermal Scrapings from Cutaneous Leishmaniasis Lesions in Tunisia. Microorganisms 2022;10:1157. [Crossref] [PubMed]
26. Stensvold CR, Høst AV, Belkessa S, et al. Evaluation of the NovaLisa™ Leishmania Infantum IgG ELISA in A Reference Diagnostic Laboratory in A Non-Endemic Country. Antibodies (Basel) 2019;8:20. [Crossref] [PubMed]
27. Bracamonte ME, Álvarez AM, Sosa AM, et al. High performance of an enzyme linked immunosorbent assay for American tegumentary leishmaniasis diagnosis with Leishmania (Viannia) braziliensis amastigotes membrane crude antigens. PLoS One 2020;15:e0232829. [Crossref] [PubMed]
28. De Silva NL, De Silva VNH, Deerasinghe ATH, et al. Validation of an In-House ELISA Method in the Diagnosis of Cutaneous Leishmaniasis Caused by Leishmania donovani in Hambantota District, Sri Lanka. Microorganisms 2022;10:921. [Crossref] [PubMed]
29. Viana SM, Montoya AL, Carvalho AM, et al. Serodiagnosis and therapeutic monitoring of New-World tegumentary leishmaniasis using synthetic type-2 glycoinositolphospholipid-based neoglycoproteins. Emerg Microbes Infect 2022;11:2147-59. [Crossref] [PubMed]
30. Schallig HDFH, Hu RVP, Kent AD, et al. Evaluation of point of care tests for the diagnosis of cutaneous leishmaniasis in Suriname. BMC Infect Dis 2019;19:25. [Crossref] [PubMed]
31. Bennis I, Verdonck K, El Khalfaoui N, et al. Accuracy of a Rapid Diagnostic Test Based on Antigen Detection for the Diagnosis of Cutaneous Leishmaniasis in Patients with Suggestive Skin Lesions in Morocco. Am J Trop Med Hyg 2018;99:716-22. [Crossref] [PubMed]
32. Jalali H, Enayati AA, Fakhar M, et al. Reemergence of zoonotic cutaneous leishmaniasis in an endemic focus, northeastern Iran. Parasite Epidemiol Control 2021;13:e00206. [Crossref] [PubMed]
33. Asfaram S, Fakhar M, Mirani N, et al. HRM-PCR is an accurate and sensitive technique for the diagnosis of cutaneous leishmaniasis as compared with conventional PCR. Acta Parasitol 2020;65:310-6. [Crossref] [PubMed]
34. Veasey JV, Zampieri RA, Lellis RF, et al. Identification of Leishmania species by high-resolution DNA dissociation in cases of American cutaneous leishmaniasis. An Bras Dermatol 2020;95:459-68. [Crossref] [PubMed]
35. Morais RCS, Melo MGN, Goes TC, et al. Duplex qPCR for Leishmania species identification using lesion imprint on filter paper. Exp Parasitol 2020;219:108019. [Crossref] [PubMed]
36. Nateghi Rostami M, Darzi F, Farahmand M, et al. Performance of a universal PCR assay to identify different Leishmania species causative of Old World cutaneous leishmaniasis. Parasit Vectors 2020;13:431. [Crossref] [PubMed]
37. Khoshnood S, Mohaghegh MA, Ghafori K, et al. A comparison of the efficiency of molecular and conventional methods in the diagnosis of cutaneous leishmaniosis. Ann Parasitol 2021;67:39-44. [PubMed]
38. Chaouch M, Aoun K, Ben Othman S, et al. Development and Assessment of Leishmania major and Leishmania tropica Specific Loop-Mediated Isothermal Amplification Assays for the Diagnosis of Cutaneous Leishmaniasis in Tunisia. Am J Trop Med Hyg 2019;101:101-7. [Crossref] [PubMed]
39. Garcia GC, Carvalho AMRS, Duarte MC, et al. Development of a chimeric protein based on a proteomic approach for the serological diagnosis of human tegumentary leishmaniasis. Appl Microbiol Biotechnol 2021;105:6805-17. [Crossref] [PubMed]
40. Yeboah C, Mosore MT, Attram N, et al. A Preliminary Study to Compare Recombinase Polymerase Amplification-Lateral Flow and Quantitative PCR in the Detection of Cutaneous Leishmania in Communities from the Volta Region of Ghana. Vector Borne Zoonotic Dis 2023;23:75-80. [Crossref] [PubMed]
41. Tayachi I, Galai Y, Ben-Abid M, et al. Use of Immunomagnetic Separation tool in Leishmania promastigotes capture. Acta Trop 2021;215:105804. [Crossref] [PubMed]
42. Kariyawasam R, Valencia BM, Lau R, et al. Evaluation of a point-of-care molecular detection device for Leishmania spp. and intercurrent fungal and mycobacterial organisms in Peruvian patients with cutaneous ulcers. Infection 2021;49:1203-11. [Crossref] [PubMed]
43. Castellanos-Gonzalez A, Cossio A, Jojoa J, et al. MiniPCR as a portable equipment for the molecular diagnosis of american cutaneous leishmaniasis. Acta Trop 2023;243:106926. [Crossref] [PubMed]
44. de Goes TC, de Morais RCS, de Melo MGN, et al. Analysis of the IGS rRNA Region and Applicability for Leishmania (V.) braziliensis Characterization. J Parasitol Res 2020;2020:8885070. [Crossref] [PubMed]
45. Chakkumpulakkal Puthan Veettil T, Duffin RN, Roy S, et al. Synchrotron-Infrared Microspectroscopy of Live Leishmania major Infected Macrophages and Isolated Promastigotes and Amastigotes. Anal Chem 2023;95:3986-95. [Crossref] [PubMed]
46. Welearegay TG, Diouani MF, Österlund L, et al. Ligand-Capped Ultrapure Metal Nanoparticle Sensors for the Detection of Cutaneous Leishmaniasis Disease in Exhaled Breath. ACS Sens 2018;3:2532-40. [Crossref] [PubMed]
47. Dueñas E, Nakamoto JA, Cabrera-Sosa L, et al. Novel CRISPR-based detection of Leishmania species. Front Microbiol 2022;13:958693. [Crossref] [PubMed]
48. Suárez M, Valencia BM, Jara M, et al. Quantification of Leishmania (Viannia) Kinetoplast DNA in Ulcers of Cutaneous Leishmaniasis Reveals Inter-site and Inter-sampling Variability in Parasite Load. PLoS Negl Trop Dis 2015;9:e0003936. [Crossref] [PubMed]
49. Sandoval Pacheco CM, Araujo Flores GV, Favero Ferreira A, et al. Histopathological features of skin lesions in patients affected by non-ulcerated or atypical cutaneous leishmaniasis in Honduras, Central America. Int J Exp Pathol 2018;99:249-57. [Crossref] [PubMed]
50. Schuster FL, Sullivan JJ. Cultivation of clinically significant hemoflagellates. Clin Microbiol Rev 2002;15:374-89. [Crossref] [PubMed]
51. Pagheh A, Fakhar M, Mesgarian F, et al. An improved microculture method for diagnosis of cutaneous leishmaniasis. J Parasit Dis 2014;38:347-51. [Crossref] [PubMed]
52. Fernandez-Flores A. A new scenario in the immunohistochemical diagnosis of cutaneous leishmaniasis. J Cutan Pathol 2017;44:1051-2. [Crossref] [PubMed]
53. Caballero ZC, Sousa OE, Marques WP, et al. Evaluation of serological tests to identify Trypanosoma cruzi infection in humans and determine cross-reactivity with Trypanosoma rangeli and Leishmania spp. Clin Vaccine Immunol 2007;14:1045-9. [Crossref] [PubMed]
54. Elmahallawy EK, Sampedro Martinez A, Rodriguez-Granger J, et al. Diagnosis of leishmaniasis. J Infect Dev Ctries 2014;8:961-72. [Crossref] [PubMed]
55. Duarte MC, Pimenta DC, Menezes-Souza D, et al. Proteins Selected in Leishmania (Viannia) braziliensis by an Immunoproteomic Approach with Potential Serodiagnosis Applications for Tegumentary Leishmaniasis. Clin Vaccine Immunol 2015;22:1187-96. [Crossref] [PubMed]
56. Menezes-Souza D, Mendes TA, Nagem RA, et al. Mapping B-cell epitopes for the peroxidoxin of Leishmania (Viannia) braziliensis and its potential for the clinical diagnosis of tegumentary and visceral leishmaniasis. PLoS One 2014;9:e99216. [Crossref] [PubMed]
57. Barbé B, Verdonck K, El-Safi S, et al. Rapid Diagnostic Tests for Neglected Infectious Diseases: Case Study Highlights Need for Customer Awareness and Postmarket Surveillance. PLoS Negl Trop Dis 2016;10:e0004655. [Crossref] [PubMed]
58. Boelaert M, Verdonck K, Menten J, et al. Rapid tests for the diagnosis of visceral leishmaniasis in patients with suspected disease. Cochrane Database Syst Rev 2014;2014:CD009135. [Crossref] [PubMed]
59. Boelaert M, Bhattacharya S, Chappuis F, et al. Evaluation of rapid diagnostic tests: visceral leishmaniasis. Nat Rev Micro 2007;5:S30-9. [Crossref]
60. Cunningham J, Hasker E, Das P, et al. A global comparative evaluation of commercial immunochromatographic rapid diagnostic tests for visceral leishmaniasis. Clin Infect Dis 2012;55:1312-9. [Crossref] [PubMed]
61. de Ruiter CM, van der Veer C, Leeflang MM, et al. Molecular tools for diagnosis of visceral leishmaniasis: systematic review and meta-analysis of diagnostic test accuracy. J Clin Microbiol 2014;52:3147-55. [Crossref] [PubMed]
62. Andrade RV, Massone C, Lucena MN, et al. The use of polymerase chain reaction to confirm diagnosis in skin biopsies consistent with American tegumentary leishmaniasis at histopathology: a study of 90 cases. An Bras Dermatol 2011;86:892-6. [Crossref] [PubMed]
63. Mousavi T, Shokohi S, Abdi J, et al. Determination of genetic diversity of Leishmania species using mini-circle kDNA, in Iran-Iraq countries border. Trop Parasitol 2018;8:77-82. [Crossref] [PubMed]
64. Cupolillo E, Grimaldi Júnior G, Momen H, et al. Intergenic region typing (IRT): a rapid molecular approach to the characterization and evolution of Leishmania. Mol Biochem Parasitol 1995;73:145-55. [Crossref] [PubMed]
65. Mugasa CM, Laurent T, Schoone GJ, et al. Simplified molecular detection of Leishmania parasites in various clinical samples from patients with leishmaniasis. Parasit Vectors 2010;3:13. [Crossref] [PubMed]
66. Cataldo JI, de Queiroz Mello FC, Mouta-Confort E, et al. Immunoenzymatic assay for the diagnosis of American tegumentary leishmaniasis using soluble and membrane-enriched fractions from infectious Leishmania (Viannia) braziliensis. J Clin Lab Anal 2010;24:289-94. [Crossref] [PubMed]
67. Cossio A, Jojoa J, Castro MDM, et al. Diagnostic performance of a Recombinant Polymerase Amplification Test-Lateral Flow (RPA-LF) for cutaneous leishmaniasis in an endemic setting of Colombia. PLoS Negl Trop Dis 2021;15:e0009291. [Crossref] [PubMed]
68. López-Vélez R, Laguna F, Alvar J, et al. Parasitic culture of buffy coat for diagnosis of visceral leishmaniasis in human immunodeficiency virus-infected patients. J Clin Microbiol 1995;33:937-9. [Crossref] [PubMed]
69. Sanger F, Air GM, Barrell BG, et al. Nucleotide sequence of bacteriophage phi X174 DNA. Nature 1977;265:687-95. [Crossref] [PubMed]
70. Osorio-Peralta DC, Petano-Duque JM, Rondón-Barragán IS. Leishmania spp. diagnosis and therapeutic management in a cat from urban area in Ibagué (Colombia). Vet Parasitol Reg Stud Reports 2024;47:100980. [Crossref] [PubMed]
71. Adams ER, Gomez MA, Scheske L, et al. Sensitive diagnosis of cutaneous leishmaniasis by lesion swab sampling coupled to qPCR. Parasitology 2014;141:1891-7. [Crossref] [PubMed]
72. Hu T, Chitnis N, Monos D, et al. Next-generation sequencing technologies: An overview. Hum Immunol 2021;82:801-11. [Crossref] [PubMed]
73. Holman HY, Martin MC, McKinney WR. Synchrotron-Based FTIR Spectromicroscopy: Cytotoxicity and Heating Considerations. J Biol Phys 2003;29:275-86. [Crossref] [PubMed]
74. Bouabid C, Yamaryo-Botté Y, Rabhi S, et al. Fatty Acid Profiles of Leishmania major Derived from Human and Rodent Hosts in Endemic Cutaneous Leishmaniasis Areas of Tunisia and Algeria. Pathogens 2022;11:92. [Crossref] [PubMed]
75. Miekisch W, Schubert JK, Noeldge-Schomburg GF. Diagnostic potential of breath analysis--focus on volatile organic compounds. Clin Chim Acta 2004;347:25-39. [Crossref] [PubMed]
76. Myhrvold C, Freije CA, Gootenberg JS, et al. Field-deployable viral diagnostics using CRISPR-Cas13. Science 2018;360:444-8. [Crossref] [PubMed]
77. Nguyen LT, Rananaware SR, Pizzano BLM, et al. Clinical validation of engineered CRISPR/Cas12a for rapid SARS-CoV-2 detection. Commun Med (Lond) 2022;2:7. [Crossref] [PubMed]