Diagnosis of leishmaniasis

Review Article Diagnosis of leishmaniasis Ehab Kotb Elmahallawy1,4, Antonio Sampedro Martínez3,Javier Rodriguez-Granger3, Yannick HoyosMallecot3, Ahmad Agil2, José Marí Navarro Mari3, José Gutierrez Fernández3,4 1 Departament of Zoonotic Diseases, Faculty of Veterinary Medicine, Sohag University, Sohag, Egypt 2 Department of Pharmacology, Faculty of Medicine, University of Granada, Granada, Spain 3 Service of Microbiology, University Hospital Virgen de las Nieves, Granada, Spain 4 Department of Microbiology, Faculty of Medicine, University of Granada, Granada, Spain Abstract Leishmaniasis is a clinically heterogeneous syndrome caused by intracellular protozoan parasites of the genus Leishmania. The clinical spectrum of leishmaniasis encompasses subclinical ( not apparent), localized (skin lesion), and disseminated (cutaneous, mucocutaneous, and visceral) infection. This spectrum of manifestations depends on the immune status of the host, on the parasite, and on immunoinflammatory responses. Visceral leishmaniasis causes high morbidity and mortality in the developing world. Reliable laboratory methods become mandatory for accurate diagnosis, especially in immunocompromised patients such as those infected with HIV. In this article, we review the current state of the diagnostic tools for leishmaniasis, especially the serological test. Key words: leishmaniasis; diagnosis; serology; HIV co-infection. J Infect Dev Ctries 2014; 8(8):961-972. doi:10.3855/jidc.4310 (Received 06 October 2013 – Accepted 27 May 2014) Copyright © 2014 Elmahallawy et al. This is an open-access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. Introduction The term leishmaniasis refers to a group of vectorborne diseases caused by obligate intracellular protozoan parasites of the genus Leishmania, belonging to the family Trypanasomatidae, order Kinetoplastidia. Natural transmission of the parasite occurs primarily via the bite of infected female sandflies of the genus Phlebotomous in the Old World and Lutzomyia in the New World. The disease can also be transmitted by shared syringes among intravenous drug abusers [1], by blood transfusion [2], by venereal infections [3,4], and congenitally from mother to infant [5]. However, these modes of transmission are very rare compared with vector-borne transmission. In a domestic environment, dogs are the most important reservoirs, maintaining the parasite endemic focus. Rodents also play a role as Leishmania reservoirs [68]. Leishmaniasis occurs in three clinical forms: (i) cutaneous leishmaniasis (CL), which is caused by L. major, L. tropica, L. aethiopica (Old World CL), L. infantun, L. chagasi (Mediterranean and Caspian Sea region CL), L. amazonensis, L. mexicana, L. braziliensis, L. panamesis, L. peruviana, and L. guayanensis (New World CL); (ii) mucocutaneous leishmaniasis (MCL) or espundia, which is caused by L. brazilensis, L. panamensis, L. guyanensis in the New World and is occasionally encountered in the Old World, caused by L. infantum and L. donovani; (iii) visceral leishmaniasis (VL), which is caused by species of the L. donovani complex that consist mainly of L. infantum, L. donovani, and L. chagasi. LV is also known as kala-azar, black fever, and Dumdum fever. There is a fourth form, known as diffuse cutaneous leishmaniasis (DCL), which is caused by L. amazonensis and L. aethiopica [9]. CL is the most prevalent form and manifests as skin lesions that can heal spontaneously. VL is the most severe form of the disease and, left untreated, is usually fatal. Post-kala-azar dermal leishmaniasis (PKDL) is a complication of LV, characterized by a macular, maculo-papular, or nodular rash on the face, torso, or other part of the body. It occurs mainly in East Africa and the Indian subcontinent, behaving as the major reservoir of the parasite in areas of anthroponotic transmission [10]. The mucocutaneous form causes partial or total mutilation of mucous membranes in the nose, mouth, and throat. Leishmania infections are worldwide in distribution; they have been reported in 98 countries Elmahallawy et al. – Diagnosis of leishmaniasis J Infect Dev Ctries 2014; 8(8):961-972. 962 on all continents. The disease is endemic in tropical and subtropical regions. There are an estimated 1.3 million new cases worldwide annually, of which 1 million are CL or MCL and another 300,000 cases are VL, with approximately 200 million people at risk of infection [11]. Of the 1.3 million estimated cases, only 600,000 are actually reported [12]. VL occurs mainly (90% of cases) in Bangladesh, Brazil, Ethiopia, India, Nepal, South Sudan, and Sudan. The governments of these countries have committed to eliminate VL by 2015 and aim to reduce the incidence of VL to 1 per 10,000 population in endemic districts [13]. VL is an important opportunistic infection in AIDS patients; in countries where there is poor access to antiretroviral therapy, the prevalence of visceral disease is rising [14]. These alarming facts have been attributed in part to the absence of an effective VL vaccine [15]. CL is found primarily in Afghanistan, Algeria, Brazil, Colombia, the Islamic Republic of Iran, Pakistan, Peru, Saudi Arabia, the Syrian Arab Republic, and Tunisia. Almost 90% of MCL cases occur in Brazil, Peru, and Bolivia. The health burden of CL in these countries remains largely unknown, partly because those who are most affected live in remote areas and often do not seek medical attention [12,15]. Control of leishmaniasis requires a combination of intervention strategies; early diagnosis and treatment is an important aspect. In VL, diagnosis is made by combining clinical signs with parasitological or serological tests (rapid diagnostic tests and others). In addition, an accurate diagnostic test that can identify active VL versus asymptomatic disease remains a key component of measures that aim to control this serious disease [16]. In CL and MCL, serological tests have limited value. Table 1 presents a summary of all the serological and antigen detection techniques commonly used for leishmaniasis diagnosis. Diagnosis of cutaneous and mucocutaneous leishmaniasis In the laboratory, diagnosis is made microscopically by direct identification of amastigotes in Giemsa-stained lesion smears of biopsies, scrapings, or impression smears. Amastigotes are observed as round or oval bodies, 2–4 μm in diameter, with characteristic nuclei and kinetoplasts. The material from the ulcer base usually has the highest yield; however, other authors have not found significant differences in the diagnostic outcomes when smears or culture samples are taken from the center or the border of the ulcer or from an incision made tangential from the ulcer [17,18]. Conventionally, three to five aspirates from different lesions or portions of lesions are dissected. The first samples should be used for microscopy and the last for culture to minimize the risk of contamination [19]. A combination of microscopy and culture increases diagnostic sensitivity to more than 85% [18,20]. Antileishmanial antibodies can be detected by serological tests; however, these are not the usual methods, because antibodies tend to be undetectable or present in low titers due to poor humoral response [2123]. Culture and DNA detection by PCR are sensitive but are not currently practical in developing countries. Diagnosis of visceral leishmaniasis In developing countries where the disease is not prevalent, the existence of laboratory facilities enables an adequate and efficient follow-up of the disease. However, in developing countries with large numbers of patients in rural areas, simple diagnostic tools are necessary for field use [24]. Laboratory diagnosis of VL includes microscopic observation and culture from adequate samples, antigen detection, serological tests, and detection of parasite DNA. Culture and microscopic observation Definitive VL diagnosis is supported by direct demonstration of parasites in clinical specimens and specific molecular methods [25-27]. The commonly used samples are splenic or bone marrow aspirates. The presence of amastigotes can also be determined in other samples such as liver biopsies, lymph nodes, and buffy coats of peripheral blood. The sensitivity of the bone marrow stained with Giemsa is about 60% to 85%. In splenic aspirates, the sensitivity is higher (93%) [28], but sampling is associated with a risk of fatal hemorrhage in inexperienced hands. To increase the sensitivity, fluorescent dye-conjugated antibodies can be used [29]. The sensitivity in peripheral blood smears is low, especially in individuals with low parasitemia. In addition, results are dependent on technical expertise and the quality of prepared slides. Culture of the parasite can improve diagnostic sensitivity, but is tedious, time-consuming, and expensive, and thus seldom used for clinical diagnosis. There are new culture methods that improve sensitivity, such as the micro-culture method (MCM); recent modifications of this method involve using the buffy coat and peripheral blood mononuclear cells [30,31]. Elmahallawy et al. – Diagnosis of leishmaniasis J Infect Dev Ctries 2014; 8(8):961-972. 963 The culture media used may be biphasic and may include Novy-MacNeal-Nicolle medium and Tobie’s medium (conversion of amastigotes to promastigotes and monophasic medium), Schneider's insect medium, M199, and Grace's insect medium (amplifying parasite number) [29]. Antigen detection in urine Several studies have demonstrated leishmanial antigen in the urine of VL patients. In 1995, De Colmenares et al. reported two polypeptide fractions of 72–75 kDa and 123 kDa in the urine of kala-azar patients [32]. In 2002, Sarkari et al. described a urinary 5–20 kDa carbohydrate-based, heat stable antigen of VL patients [33]. A latex agglutination test (KAtex, Kalon Biological, UK) for detection of this antigen in urine samples was evaluated using samples from confirmed cases and controls from endemic and non-endemic regions. This test showed good specificity (82% to 100%), but had low to moderate sensitivity that ranged from 47% to 95% [34-38]. Nowadays, this method is useful for the diagnosis of disease in cases with deficient antibody production. In this respect, this method has reported 100% sensitivity and 96% specificity in immunocompromised patients [39]. In another study, 87% specificity and 85% sensitivity were obtained for primary VL in HIV-coinfected patients, and the method had predictive capabilities in the follow-up of treatment and detection of subclinical infection in Leishmania/HIV co-infected Table 1. Serological and antigen detection techniques commonly used for leishmaniasis diagnosis Assay or method Sensitivity (%) Specificity (%) Advantages Disadvantages References Antibody detection IFA test 87–100 77–100 Positive in the early stages of infection and undetectable six to nine months after cure. Requires sophisticated laboratory. No application in the field. 41,42,43 ELISA Can be used to screen a large scale of epidemiologic studies. Sensitivity and specificity is greatly influenced by the antigen used. Requires skilled personnel, sophisticated equipment, and electricity. CSA as antigen 80–100 84–95 45,46,47 with fucose-mannose ligand 95–100 95 53 rK39 as antigen 75–98 79–89 58,59 Western blot 90–98 98–100 Provides detailed antibody responses to various leishmanial antigens. Time consuming, technically cumbersome, and expensive. 69, 70, 71, 72, 73 DAT 85–100 91–100 Rapid test, applicable in the field. Limited to use in regions of endemicity. Long incubation time is needed. Unavailability of commercial source of the postive antigen and fragility of is aqueous form. 74,75,76,77,79 IC test with rK39 90–100 93–100 Inexpensive, rapid, simple, and can be performed by untrained person. 84,85,86,88 Antigen detection Latex agglutination test in urine (KAtex) 79–100 60–100 Simple, easy performing. Useful in diagnosis of disease immunocompromised patients. Difficult to distinguish weakly positive from negative results and the urine must be boiled to avoid falsepositive reactions. 33,34,36,37 Elmahallawy et al. – Diagnosis of leishmaniasis J Infect Dev Ctries 2014; 8(8):961-972. 964 cases [40]. Another urinary leishmanial antigen, a lowmolecular weight, heat-stable carbohydrate has been detected in the urine of VL patients by an agglutination test with 60% to 71% sensitivity and 79% to 94% specificity [41]. In summary, the latex agglutination test is simple, easy to perform, inexpensive, rapid, and can be used as a screening test. Efforts are being made to improve the performance of this technique, because it promises to be a test of cure in populations of developing areas [28]. Serological diagnosis Specific serological diagnosis is based on the presence of a specific humoral response. Current serological tests are based on four formats: indirect fluorescent antibody (IFA), enzyme-linked immunosorbent assay (ELISA), western blot, and direct agglutination test (DAT). The sensitivity depends upon the assay and its methodology, but the specificity depends on the antigen rather than the serological format used. It has been noted that all antibody detection tests share the same drawbacks; the antibodies remain positive for many months after the patient has been cured and do not differentiate between current and past infection. In endemic regions, asymptomatically infected persons can also be positive in these tests. Indirect fluorescent antibody (IFA) test The IFA test shows acceptable sensitivity (87%– 100%) and specificity (77%–100%) [41,42]. Promastigote forms should be the antigens of choice for diagnosis of visceral leishmaniasis by the IFA test because they minimize cross-reactivity with trypanosomal sera [43]. The antibody response is detectable very early in infection and becomes undetectable six to nine months after cure; hence, if the antibodies persist in low titers, it is a good indication of a probable relapse [44]. The need for a sophisticated laboratory with a fluorescence microscope restricts use of the IFA test to reference laboratories. Enzyme-linked immunosorbent assay (ELISA) ELISA is the preferred laboratory test for serodiagnosis of VL. The technique is highly sensitive, but its specificity depends upon the antigen used. Moreover, this assay can be performed easily and is adaptable for use with several antigenic molecules. One of the antigens used in the ELISA test is a crude soluble promastigote antigen (CSA) that is obtained by freezing and thawing live promastigotes. The sensitivity of ELISA using CSA ranges from 80% to 100%, and specificity ranges from 84% to 95% [45,46]. Cross-reactivity with sera from patients with tuberculosis, trypanosomiasis, and toxoplasmosis has been reported [47-51]. When selective antigenic molecules were used (with molecular weights of 116 kDa, 72 kDa, and 66 kDa), the specificity approached 100%, while the sensitivity was very low (37%) [31,52]. Other purified antigens used were a 36-kDa glycoprotein [53], metabolic antigens released by L. donovani [54], and A2 proteins implicated in the development of visceral disease [55-57]. A conserved portion of a kinesin-related protein recombinant antigen from a cloned protein of L. chagasi, called rK39, has been reported to be highly reactive to sera from human and canine VL cases when run in an ELISA format [58,59]. Using this recombinant antigen, 99% specificity and sensitivity were reported in immunocompetent patients with clinical VL. In India, this antigen was reported with a sensitivity of 98% and a specificity of 89% [60]; however, a report from Sudan and other countries revealed that this antigen showed low sensitivity (75%) and specificity (70%) [61]. In HIV-positive patients, rK39-ELISA showed higher sensitivity (82%) than the IFA test (54%), with higher predictive value for detecting VL [62]. With successful therapy, antibody titers declined steeply at the end of treatment and during follow-up; in contrast, patients who relapsed showed increased titers of antibodies to rK39. This suggests the possible application of rK39-ELISA in monitoring drug therapy and detecting relapse of VL [51]. In another study, two hydrophilic antigens of Leishmania chagasi were used (rk9 and rk26), leading to an increase in the list of available antigens for serodiagnosis of VL [63]. Another kinesin recombinant related protein used in ELISA assay is rKE16. The use of this antigen for VL diagnosis has been very sensitive and specific as rK39 when tested in patients from China, Pakistan, and Turkey [64]. Another new assay, based on the detection of the K28 fusion protein in studies performed in Sudan and Bangladesh with 96% sensitivity in Sudan and 98% in Bangladesh, has been developed [65]. Moreover, heat shock proteins HPS70 or histones proteins H2A, H2B, H3, and H4 may have potential use for serodiagnosis of VL [66,67]; furthermore, lipid-binding proteins (LBPs) as antigens have shown high levels of sensitivity and an absence of cross-reactions with the sera of patients with other diseases [69]. The ELISA test, due to the requirement of skilled personnel, Elmahallawy et al. – Diagnosis of leishmaniasis J Infect Dev Ctries 2014; 8(8):961-972. 965 sophisticated equipment, and electricity, is not used in endemic regions for the diagnosis of VL. Immunoblotting (western blot) For this type of testing, promastigotes are cultured to log phase, lysed, and the proteins are separated on SDS-PAGE. The separated proteins are electrotransferred onto a nitrocellulose membrane and probed with serum from the patient. The western blot technique provides detailed antibody responses to various leishmanial antigens [69,70], and has been found to be more sensitive than the IFA test and ELISA, especially in co-infected HIV patients with VL [71-73], but the drawbacks of the technique (equipment and time requirements, cumbersomeness, and cost) limit its use to research laboratories. Direct agglutination test (DAT) The DAT is based on direct agglutination of Leishmania promastigotes that react specifically with anti-Leishmania antibodies in the serum specimen, resulting in agglutination of the promastigotes. Whole, trypsinized, coomassie-stained promastigotes can be used either as a suspension or in freeze-dried form that can be stored at room temperature for at least two years, facilitating its use in the field [74,75]. Chappuis et al. [76], in a meta-analysis that included thirty studies evaluating DAT, found that the DAT had sensitivity and specificity estimates of 94.8% (95% confidence intervals [CI], 92.7–96.4) and 97.1% (95% CI, 93.9–98.7), respectively. Moreover, in settings where parasitological confirmation is not feasible, the freeze-dried DAT together with classical clinical features of VL can be used for diagnosis at a cut-off for positive DAT, which is 1:12,800 as in endemic areas. The DAT is simpler than many other tests but presents severe problems in terms of reproducibility of results, which depends on antigen elaboration [77]. A new antigen elaboration method, the EasyDAT described in 2003, shows the same sensitivity, specificity, and durability as the traditional DAT antigen method but offers the additional advantages of cost reduction and standardization [78]. Although the DAT for the serodiagnosis of visceral leishmaniasis has high sensitivity and specificity, it still has some limitations; among these are the relative long incubation time (18 hours) and the serial dilutions of the samples that must be made. In order to circumvent these problems, Schoone et al. in 2001 [79] developed a fast agglutination screening test (FAST). The FAST utilizes only one serum dilution (qualitative result) and requires three hours of incubation. This makes the test very suitable for the screening of large populations. The sensitivity and specificity of FAST were found to be 91.1%–95.4% and 70.5%–88.5%, respectively [80,81]. Anti-Leishmania antibodies may persist for years as a result of previous VL infection, so titers measured by DAT may remain positive for up to five years after recovery in > 50% of VL patients, which may limit the DAT’s widespread applicability in regions of endemicity. Although the DAT is the first real field test, it remains the serological test of choice as well as the first antibody detection test for VL used in field settings, particularly in many developing countries and in Leishmania/HIV co-infections [73,82,83]. Immunochromatographic assay (IC) The immunochromatographic test using rK39 antigen (39 amino-acid-repeat recombinant leishmanial antigen from L. chagasi) has become popular in recent years. It is a qualitative membranebased immunoassay with nitrocellulose strips impregnated with recombinant K39 Leishmania antigen. A drop of blood or serum is smeared over the pad of the strips and dipped in a small amount of buffer; the results are ready within a few minutes. In clinical cases of VL, the rK39 IC showed variation in the sensitivity and specificity among different populations. The rK39 IC showed 100% sensitivity and 93%–98% specificity in India [84,85], 90% sensitivity and 100% specificity in Brazil [86], and 100% sensitivity and specificity in the Mediterranean area [87]. In other reports in southern Europe, the rK39 IC test was positive in only 71.4% of the cases of VL [88]; in Sudan, rK39 IC showed a sensitivity of 67% [61]. These differences in sensitivity may be due to differences in the antibody responses observed in different ethnic groups. The rK39 IC assay has proven to be versatile in predicting acute infection, and it is the only available format for diagnosis of VL with acceptable sensitivity and specificity levels. It is also easy to use in the field, rapid (15–20 minutes), cheap, and gives reproducible results. Like the DAT assay, IC is positive in a significant proportion of healthy individuals in endemic regions and for long periods after cure; hence, this limits its usefulness in persons with a previous history of VL who present with recurrence of fever and splenomegaly, as these tests cannot discriminate between a case of VL relapse and other pathologies. Elmahallawy et al. – Diagnosis of leishmaniasis J Infect Dev Ctries 2014; 8(8):961-972. 966 Latex agglutination test (LAT) The LAT is one of the recently developed rapid diagnostic tests for the rapid detection of antiLeishmania antibodies against the A2 antigen derived from the amastigote form as well as those against crude antigens derived from the promastigote form of an Iranian strain of L. infantum. In a comparative study with the DAT, the sensitivity of tested human sera from DAT-confirmed patients yielded 88.4% sensitivity, while the specificity was 93.5% on A2LAT amastigote, with a higher degree of similarity in accuracy to the DAT [89,90]. Molecular methods: polymerase chain reaction (PCR) Although different molecular methods have successively been evaluated for leishmaniasis diagnosis, PCR-based assays are the main molecular diagnostic tools, especially in immunosupressed patients [91-94]. PCR protocols to detect Leishmania DNA in VL diagnosis have used a variety of samples, including spleen, lymph node, and bone marrow aspirates, whole blood, and buffy coat [92,95-101]. There are different target sequences used, which include ribosomal RNA genes, kinetoplast DNA (kDNA), miniexon-derived RNA (medRNA), and the β-tubulina gene region [103]. Many PCR-based methods for diagnosis of VL have been described with different specificities and sensitivities; PCR assay sensitivity depends on the sample used. Sensitivity is highest (near 100%) in spleen or bone marrow [93,97,102] samples. The ideal sample is peripheral blood due to its non-invasive character. Using peripheral blood, the sensitivity ranges described vary from 70% to 100% [93,102,103]. A comparative clinical study between conventional microbiologic techniques and a leishmania speciesspecific PCR assay in HIV-co-infected and HIVuninfected patients has shown the sensitivity of the leishmania species-specific PCR to be 95.7% for bone marrow and 98.5% for peripheral blood samples; the sensitivity in HIV-co-infected and non-HIV-coinfected adults was 100% [94]. A PCR-ELISA was used to diagnose VL in HIVnegative patients; peripheral blood was used and yielded a high sensitivity [104]. A similar PCR-based technique was applied in the diagnosis of VL in HIV patients, with good results [28]. Recently, an evaluation of an oligochromatography-PCR for diagnosis of VL, cutaneous leishmaniasis (CL), and post kala-azar dermal leishmaniasis (PKDL), showed a high sensitivity (> 95%) on lymph, blood, and bone marrow samples from confirmed VL patients [105]. Another interesting approach is a rapid fluorogenic PCR technique. Wortmann et al. used a fluorescent DNA probe for a conserved rRNA gene that is amplified using flanking primers; this technique using clinical samples showed great sensitivity and specificity [98]. The real-time PCR has the advantage of being quantitative, which could be useful in the follow-up of treatment, allowing for the assessment of the parasite burden [106,107]. However, PCR techniques remain complex and expensive, and in most VL-endemic countries, they are restricted to a few teaching hospitals and research centers. Diagnosis of VL-HIV co-infection According to the World Health Organization (WHO) [108], an estimated 35 million people worldwide are living with HIV. Immunosuppression may reactivate latent Leishmania infection in asymptomatic patients and among HIV/AIDS patients. It is known that Leishmania has emerged as an opportunistic disease among HIV patients in endemic areas [109-112]. Moreover, it has been noted that the risks of clinical VL in HIV patients increased by 100 to 1,000) [111]. Although cases of co-infection have so far been reported in 33 countries worldwide, most of the cases have been found in sub-Saharan African countries, especially in East Africa. In Humera (in northwest Ethiopia), the proportion of VL patients coinfected with HIV increased from 18.5% in 1998–99 to 40% in 2006 [113]. VL in HIV patients has atypical clinical presentation; only 75% of HIV-infected patients, as opposed to 95% of non-HIV-infected patients, exhibit the characteristic clinical pattern – namely, fever, splenomegaly, hepatomegaly, and gastrointestinal involvement [114-117]. The diagnostic principles remain the same as those for non-HIV-infected patients. The presence of Leishmania amastigotes in the bone marrow can often be demonstrated in buffy coat preparation and in unusual locations (stomach, colon, or lungs) [118], but it has lower sensitivity in VL-HIV patients. For HIV patients, the sensitivity of antibody-based immunological tests such as the IFA test and ELISA is low. Serological tests have limited diagnostic value because over 40% of co-infected individuals have no detectable specific antibody levels against Leishmania [119]. In their meta-analysis, Cota et al. [73] Elmahallawy et al. – Diagnosis of leishmaniasis J Infect Dev Ctries 2014; 8(8):961-972. 967 summarized the accuracy of different serological techniques used for diagnosing HIV-co-infected persons. The estimated sensitivities using random effect models and their respective 95% confidence intervals for the other tests were: IFA test, 51%; ELISA, 66% (40% to 88%); DAT, 81% (61% to 95%); and immunoblotting, 84% (75% to 91%). The estimated specificity using random effect models and their respective confidence intervals for the following tests were: immunoblotting, 82% (65% to 94%); ELISA, 90% (77% to 98%); IFA test, 93% (81% to 99%); and DAT, 90% (66% to 100%). Thus, due to the low sensitivity of the serological tests for VL diagnosis in HIV-infected patients, at least two different serological tests should be used for each patient to increase the sensitivity of antibody detection [120]. The detection of polypeptide fractions of 72–75 kDa and 123 kDa of Leishmania antigen in the urine of patients with VL was 96% sensitive and 100% specific; furthermore, these antigens were not detectable after three weeks of treatment, suggesting a good prognostic value [32]. In conclusion, serology should not be used to rule out a diagnosis of VL among HIV-infected patients; an additional specially recommended serological test and/or molecular or parasitological methods may be necessary if the results of serological tests are negative. Conclusions Leishmaniasis is a global health problem and a major killer in endemic countries. Recent years have witnessed extraordinary potential progress in diagnosing Leishmania infection. The main challenge in these trials has been to reach the gold standard test in order to establish effective strategic programs to control and eradicate the disease. There are several methods of laboratory diagnosis of leishmaniasis; these include parasite detection by microscopic examination, culture, or molecular biology-based assays for detecting parasite DNA (PCR). The molecular tests are more sensitive than microscopic examination and parasite culture, but they remain restricted to referral hospitals and research centers. It could be concluded from previous studies that serological diagnosis is an alternative tool to the invasive methods of parasitological diagnosis for large-scale and decentralized diagnosis of leishmaniasis, especially VL. These serodiagnostic techniques are, however, of limited use for CL and MCL because of low sensitivity and variable specificity. Serologic diagnosis of VL can be accomplished with many techniques, such as IFA, ELISA, western blot, rapid strip testing for rK39 antigen (IC), DAT, or detection of Leishmania antigen in urine by latex agglutination (KAtex). Selection of the serological test during diagnosis should be based on different parameters such as region, cost, sensitivity, specificity, feasibility, sustainability, and field applicability, especially in problematic endemic areas. Detection of antibody responses to parasite antigens has inherent limitations. First, high serum antibody levels are present in both asymptomatic and active VL. Second, serum antiLeishmania antibodies remain present for several years and complicate the diagnosis of relapse. Third, the specificity of these tests in VL-endemic areas is variable because there are a number of individuals with no clinical VL who have antileishmanial antibodies, complicating this specificity. Additional challenges originate from Leishmania/HIV coinfection, as the co-infection may appear without the typical clinical signs. The immunosuppressive action of Leishmania/HIV co-infection makes the serological approach to diagnosis especially challenging. However, DAT, immunoblotting, and the latex agglutination test (KAtex) have been proven to be superior in diagnosis of these co-infection cases. Negative results of serology should not reliably exclude a diagnosis of VL among HIV-infected patients. An additional molecular or other serological or parasitological test may be necessary to reach an accurate diagnosis if the results of serological tests are negative. Acknowledgements This work was supported by EK Elmahallawy, who has a PhD scholarship (number 736) from Erasmus Mundus Scholarship Programme (ELEMENT Action 1 First call). References 1. Cruz I, Morales MA, Noguer I, Rodríquez A, Alvar J (2002) Leishmania in discarded syringes from intravenous drug users. Lancet 359: 1124-1125. 2. Owens SD, Oakley DA, Marryott K, Hatchett W, Walton R, Nolan TJ, Newton A, Steurer F, Schantz P, Giger U (2001) Transmission of visceral leishmaniosis through blood transfusion from infected English foxhounds to anemic dogs. J Am Vet Med Assoc 219: 1076-1083. 3. Symmers WS (1960) Leishmaniasis acquired by contagion: A case of marital infection in Britain. Lancet 7116: 127-132. 4. Chatterjee KD (1980) Dermal leishmanoid. In: ChatterjeeKD Editor Parasitology, 12th edition. Calcutta: Chatterjee Medical Publishers. 64-65. 5. Rosypal AC, Troy GC, Zajac AM, Frank G, Lindsay DS (2005) Transplacental transmission of a North American isolate of Leishmania infantum in an experimentally infected beagle. J Parasitol 91: 970-972. Elmahallawy et al. – Diagnosis of leishmaniasis J Infect Dev Ctries 2014; 8(8):961-972. 968 6. World Health Organization (1984) The Leishmaniases, Report of a WHO Expert Committee. Technical Report Series 701 Geneva: WHO. 7. Ready PD (2008) Leishmaniasis emergence & climate change. Rev Sci Tech 27: 399-412. 8. Zijlstra EE, Musa AM, Khalil EA, el-Hassan IM, el-Hassan AM (2003) Post-kalaazar dermal leishmaniasis. Lancet Infect Dis 3 :87-98. 9. Singh S (2006) New developments in diagnosis of leishmaniasis. Indian J Med Res 123: 311-330. 10. Thakur, C P, Kumar K (1992) Post kala-azar dermal leishmaniasis: a neglected aspect of kala-azar control programmes. Ann Trop Med Parasitol 86: 355-359. 11. World Health Organization (2014). Leishmaniasis: the disease and its epidemiology Available:http:/who.int/leishmaniasis/disease_epidemiology/ en/index.htmlAccessed May 2014. 12. Alvar J, Vélez ID, Bern C, Herrero M, Desjeux P, Cano J, Jannin J, den Boer M (2012). Leishmaniasis worldwide and global estimates of its incidence. PLoS One 7: e35671. doi 10.1371/journal.pone.0035671. 13. World Health Organization (2011). Elimination visceral leishmaniasis: a multiplepronged approach. Available: http://www.who.int/tdr/news/2011/vl-elimination/en/. Accessed May 2014. 14. Alvar J, Aparicio P, Aseffa A, Den Boer M, Cañavate C, Dedet JP, Gradoni L, Ter Horst R, López-Vélez R, Moreno (2008): The relationship between leishmaniasis and AIDS: the second 10 years. Clin Microbiol Rev 21: 334-359. 15. World Health Organization (2010) Control of the leishmaniasis: report of a meeting of the WHO Expert Committee on the Control of Leishmaniases. World Health Organ Tech Rep Ser 949: 1-186. 16. Chappuis F, Sundar S, Hailu A, Ghalib H, Rijal S, Peeling RW, Alvar J, Boelaert M (2007) Visceral leishmaniasis: what are the needs for diagnosis, treatment and control? Nat Rev Microbiol 5: 873-882. 17. Weina PJ, Neafle RC, Wortmamm G, Polhemus M, Aronson NE. (2004) Old World leishmaniasis: an emerging infection among deployed US military and civilian workers. Clin Infect Dis 39: 1674-1680. 18. Ramirez JR, Adugelo S, Muskus C (2000) Diagnosis of cutaneous leishmaniasis in Colombia: the sampling site within lesions influences the sensititivity of parasitological diagnosis. J Clin Microbiol 38: 3768-3773. 19. Vega-López F (2003) Review: Diagnosis of cutaneous leishmaniasis. Curr Opin Infect Dis 16: 97-101. 20. Blum J, Desjeux P, Schwartz E, Beck B, Hazt C (2004) Treatament of cutaneous leishmaniasis among travellers. J Antimicrob Chemother 53: 158-166. 21. Romero GA, de la Gloria Orge Orge M, de Farias MV, Paes MG, de Olivera Macedo V, de Carvalho EM (2005) Antibody response in patients with cutaneous leishmaniasis infected by Lesihmania (Viannia) brazilensis or Leishmania (Viannia) guayanensis in Brazil. Acta Trop 93: 49-56. 22. Amaral V, Pirmez C Goncalves A, Ferreira V, Grimaldi G Jr (2000). Cell populations in lesions of cutaneous leishmaniasis of Leishmania amazonensis – infected rhesus macaques, Macaca mulatto. Mem Inst Oswaldo Cruz 95: 209-216. 23. Ahluwalia S, Lawn SD, Kanagalingen J, Grant H, Lockwood A (2004) Mucocutaneous leishmaniasis: an imported infection among travellers to central and South America. BMJ 329: 842-844. 24. Santarém N, Cordeiro da Silva A (2007). Diagnosis of visceral leishmaniasis. In: Mendez-Vilas A, editor. Communicating current research and educational topics and trends in applied microbiology. Badajoz: Formatex..Pag 839846. 25. Reus M, García B, Vázquez V, Morales DD, Fuster M, Sola J (2005) Visceral leishmaniasis: diagnosis by ultrasound.guided fine needle aspiration of an axillary node. Br J Radiol 78: 158-160. 26. Artan R, Yilmaz A, Akcam M, Aksoy NH (2006) Liver biopsy in the diagnosis of visceral leishmaniasis. J Gastroenterol Hepatol 21: 299-302. 27. Del Olmo ML, Aller de a Fuente R, Velayos Jimenez B, Fernandez Salazar L, Gonzalez Hernandez JM (2009) Visceral leishmaniasis diagnosed by duodenal biopsy. Rev Esp Enferm 101: 439-440. 28. Srivastava P, Dayama A, Mehrotra S, Sundar S (2011) Diagnosis of visceral leishmaniasis. Trans R Soc Trop Med Hyg 105: 1-6. 29. Zijlstra EE, Ali MS, el-Hassan AM, el-Toum IA, Satti M, Ghalib HW, Kager PA (1992) Kala-azar: a comparative study of parasitological methods and the direct agglutination test in diagnosis. Trans R Soc Trop Med Hyg 86: 505-507. 30. Allahverdiyev AM, Bagirova M, Uzun S, Alabaz D, Aksaray N, Kocabas E, Koksal F (2005) The value of a new microculture method for diagnosis of visceral leishmaniasis by using bone marrow and peripheral blood. Am J Trop Med Hyg 73: 276-280. 31. Maurya R, Mehrotra S, Prajapati VK, Nylén S, Sacks D, Sundar S (2010) Evaluation of blood agar microtiter plates for culturing leishmania parasites to titrate parasite burden in spleen and peripheral blood of patients with visceral leishmaniasis. J Clin Microbiol 48: 1932-1934. 32. De Colmenares M, Portus M, Riera C, Gallego M, Aisa MJ, Torras S (1995) Detection of 72-75-kD and 123-kD fractions of Leishmania antigen in urine of patients with visceral leishmaniasis. Am J Trop Med Hyg 52: 427-428. 33. Sarkari B, Chance M, Hommel M (2002) Antigenuria in visceral leishmaniasis: detection and partial characterisation of a carbohydrate antigen. Acta Trop 82: 339-348. 34. Attar ZJ, Chance ML, el-Safi S, Carney J, Azazy A, El-Hadi M, Dourado C, Hommel M (2001) Latex agglutination test for the detection of urinary antigens in visceral leishmaniasis. Acta Trop 78: 11-16. 35. El-Safi SH, Abdel-Haleem A, Hammad A , El Basha I, Omer A, Kareem HG, Boalert M, Chance M, Hommel M (2003). Field evaluation of latex agglutination test for detecting urinary antigens in visceral leishmaniasis in Sudan. East Mediterr Health J 9(4): 844-855. 36. Rijal S, Boelaert M, Regmi S, Karki BM, Jacquet D, Singh R, Chance ML, Chappuis F, Hommel M, Desjeux P, Van der Stuyft P, Le Ray D, Koirala S (2004) Evaluation of a urinary antigen-based latex agglutination test in the diagnosis of kala-azar in eastern Nepal 9: 724-729. 37. Malaei S, Mohebali M, Akhoundi B, Zarei Z (2006) Evaluation of latex agglutination test (Katex) for the detection of urinary antigens in human visceral leishmaniasis. J Tehran Un Med Sci 4: 101-108. 38. Diro E, Techane Y, Tefera T (2007) Field evaluation of FDDAT rK39 dipstick and KATEX (urine latex agglutination) Elmahallawy et al. – Diagnosis of leishmaniasis J Infect Dev Ctries 2014; 8(8):961-972. 969 for diagnosis of visceral leishmaniasis in northwest Ethiopia. Trans R Soc Trop Med Hyg 101: 908-914. 39. Vilaplana C, Blanco S, Domínguez J, Giménez M, Ausina V, Tural C, Muñoz C (2004) Noninvasive method for diagnosis of visceral leishmaniasis by a latex agglutination test for detection of antigens in urine samples. J Clin Microbiol 42: 1853-1854. 40. Boelaert M, El-Safi S, Hailu A, Mukhtar M, Rijal S, Sundar S, Wasunna M, Aseffa A, Mbui J, Menten J, Desjeux P, Peeling RW (2008) Diagnostic tests for Kala-azar: a multicentre study of the freeze-dried DAT rK39 strip test and KAtex in East Africa, the Indian subcontinent. Trans R Soc Trop Med Hyg 102: 32-40. 41. Boelaert M, Rijal S, Regmi S, Singh R, Karki B, Jacquet D, Chappuis F, Campino L, Desjeux P, Le Ray D, Koirala S, Van der Stuyft P (2004) A comparative study of the effectiveness of diagnostic tests for visceral leishmaniasis. Am J Trop Med Hyg 70: 72-77. 42. Iqbal J, Hira PR, Saroj G, Philip R, Al-Ali F, Madda PJ, Sher A (2002) Imported visceral leishmaniasis: diagnostic dilemmas and comparative analysis of three assays. J Clin Microbiol 40: 475-479. 43. Badaró R, Reed SG, Carvalho EM (1983) Immunofluorescent antibody test in American visceral leishmaniasis: sensitivity and specificity of different morphological forms of two Leishmania species. Am J Trop Med Hyg 32: 480-484. 44. 44.Singh S, Sivakumar R (2003) Recent advances in the diagnosis of leishmaniasis. J Postgrad Med 49: 55-60. 45. Ryan JR, Smithyman AM, Rajasekariah GH, Hochberg L, Stiteler JM, MartinSK (2002) Enzyme-linked immunosorbent assay based on soluble promastigote antigen detects immunoglobulin M (IgM) & IgG antibodies in sera from cases of visceral and cutaneous leishmaniasis. J Clin Microbiol 40: 1037-1043. 46. Rajasekariah GH, Ryan JR, Hillier SR, Yi LP, Stiteler JM, Cui L, Smithyman AM, Martin SK (2001) Optimisation of an ELISA for the serodiagnosis of visceral leishmaniasis using in vitro derived promastigote antigens. J Immunol Methods 252: 105-119. 47. Bray RS (1976) Immunodiagnosis of leishmaniasis. In Cohen S, Sadun EH, editors. Immunology of parasitic infections. Oxford: Blackwell. 65-76. 48. Smrkovski LL, Larson CL (1977) Antigenic cross reactivity between Mycobacterium bovis (BCG) and Leishmania donovani. Infect Immun 18: 561-562. 49. 49 Bray RS (1985) Immunodiagnosis of leishmaniasis. In Chang KP, Bray RS, editors. Leishmaniasis. Amsterdam: Elsevier. 177-181. 50. 50 Choudhry A, Guru PY, Saxena RP, Tandon A, Saxena KC (1990) Enzyme-linked immunosorbent assay in the diagnosis of kala-azar in Bhadohi (Varanasi) India. Trans R Soc Trop Med Hyg 84: 363-366. 51. Kumar R, Pai K, Pathak K, Sundar S (2001) Enzyme-linked immunosorbent assay for recombinant K39 antigen in diagnosis, prognosis of Indian visceral leishmaniasis. Clin Diagn Lab Immunol 8: 1220-1224. 52. Vinayak VK, Mahaja D, Sobt RC, Singl N, Sunda S (1994) Anti-66 kDa anti-leishmanial antibodies as specific immunodiagnostic probe for visceral leishmaniasis. Indian J Med Res 99: 109-114. 53. 53. Palatnik-de-Sousa CB, Gomes EM, Paraguai-de-Souza E, Palatnik M, Luz K, Borojevic R (1995) Leishmania donovani: titration of the antibodies to the fucose-mannose ligand as an aid in the diagnosis and prognosis of visceral leishmaniasis. Trans R Soc Trop Med Hyg 89: 390-393. 54. Martin SK, Thuita-Harun L, Adoyo-Adoyo M, Wasunna KM (1998) A diagnostic ELISA for visceral leishmaniasis based on antigen from media conditioned by Leishmania donovani promastigotes. Ann Trop Med Parasitol 92: 571-577. 55. Carvalho FA, Charest H, Tavares CA, Matlashewski G, Valente EP, Rabello A, Gazzinelli RT, Fernandes AP (2002) Diagnosis of American visceral leishmaniasis in humans and dogs using the recombinant Leishmania donovani A2 antigen. Diagn Microbiol Infect Dis 43: 289-295. 56. McCall L, Zhang W, Matlashewki G (2013) Determinants for the Development of Visceral Leishmaniasis Disease. PLOS Pathog 9: e1003053. 57. Rafati S, Hassani N, Taslimi Y, Movassagh H, Rochette A, Papadopoulou B (2006) Amastin peptide-binding antibodies as biomarkers of active human visceral leishmaniasis. Clin Vacc Immunol 13: 1104-1110. 58. Burns JM Jr, Shreffler WG, Benson DR, Ghalib HW, Badaro R, Reed SG (1993) Molecular characterization of a kinesin-related antigen of Leishmania chagasi that detects specific antibody in both African and American visceral leishmaniasis. Proc Natl Acad Sci USA 90: 775-790. 59. Badaró R, Benson D, Eulálio MC, Freire M, Cunha S, Netto EM, Pedral-Sampaio D, Madureira C, Burns JM, Houghton RL, David JR, Reed SG (1996) rK39: A Cloned Antigen of Leishmania chagasi that Predicts Active Visceral Leishmaniasis J Infect Dis 173: 758-776. 60. Singh DP, Goyal RK, Singh RK, Sundar S, Mohapatra TM (2010) In search of an ideal test for diagnosis, prognosis of kala-azar. J Health Popul Nutr 28: 281-285. 61. Zijlstra EE, Nur Y, Desjeux P, Khalil EA, El-Hassan AM, Groen J (2001) Diagnosing visceral leishmaniasis with the recombinant K39 strip test: experience from the Sudan. Trop Med Int Health 6: 108-113. 62. Houghton RL, Petrescu M, Benson DR, Skeiky YA, Scalone A, Badaró R, Reed SG, Gradoni L (1998) A cloned antigen (recombinant K39) of Leishmania chagasi diagnostic for visceral leishmaniasis in human immunodeficiency virus type 1 patients and a prognostic indicator for monitoring patients undergoing drug therapy. J Infect Dis 177: 13391344. 63. Mohapatra TM, Singh DP, Sen MR, Bharti K, Sundar S (2010) Comparative evaluation of rK9 rK26 and rK39 antigens in the serodiagnosis of Indian visceral leishmaniasis. J Infect Dev Ctries 4: 114-117. doi:10.3855/jidc.544. 64. Sivakumar R, Sharma P, Chang KP, Singh S (2006) Cloning expression, purification of a novel recombinant antigen from Leishmania donovani. Protein Expr Purif 46: 156-165. 65. Pattabhi S, Whittle J, Mohamath R, El-Safi S, Moulton GG, Guderian JA, Colombara D, Abdoon AO, Mukhtar MM, Mondal D, Esfandiari J, Kumar S, Chun P, Reed SG, Bhatia A (2010) Design development and evaluation of rK28-based point-of-care tests for improving rapid diagnosis of visceral leishmaniasis. PLoS Negl Trop Dis 4: e822. 66. Amorim AG, Carrington M, Miles MA, Barker DC, de Almeida ML (1996) Identification of the C-terminal region of 70 kDa heat shock protein from Leishmania (Viannia) braziliensis as a target for the humoral immune response. Cell Stress Chaperones 1: 177-187.