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Diagnosis of infections with Leishmania infantum using PCR–ELISA
Published online by Cambridge University Press: 12 July 2001
Abstract
On the basis of partial amplification of a cloned fragment of kDNA of Leishmania infantum which is specific for this species, we developed a PCR–ELISA technique which avoids the problems associated with classical diagnostic techniques. This technique was tested on 33 L. infantum strains from 19 different zymodemes, which were recognized equally. It was also used on human and canine clinical samples. PCR–ELISA has a higher sensitivity than the other techniques used (IFAT, parasite cultures, optical microscopy of stained samples) and permits detection of a minimum of 0.1 promastigotes or 1 fg of genomic DNA. PCR–ELISA can be used to diagnose human cutaneous leishmaniasis using material obtained by scraping the lesion margin, and human visceral leishmaniasis in HIV(+) individuals and canine leishmaniasis with peripheral blood samples. The presence of L. infantum in dogs with low antibody titres with IFAT technique (20 and 40) was demonstrated indicating that seroprevalence data from epidemiological studies underestimate the true rates of infection.
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INTRODUCTION
Leishmania infantum is the aetiological agent of leishmaniasis in Spain where it is responsible for both the cutaneous (HCL) and the visceral forms (HVL). Those include cases of coinfection with HIV, with important clinical, diagnostic, chemotherapeutic, epidemiological and economic complications (WHO, 1996). It is also responsible for canine leishmaniasis (CaL) which has been the focus of a number of studies in Spain to determine seroprevalence rates for Leishmania. These normally range from 5 to 10% but rise to above 20% in some areas (Martínez-Cruz et al. 1990; Fisa et al. 1992; Nieto et al. 1992; Acedo-Sánchez et al. 1996; Morillas et al. 1996). A similar situation has been reported for other southern European countries such as Portugal, France and Italy (Lanotte et al. 1978; Abranches et al. 1983; Bettini & Gradoni, 1986; Jambou et al. 1986; Semiao Santos et al. 1995). One very interesting aspect of the epidemiological studies of CaL is the high percentage of dogs with a titre of anti-Leishmania antibodies lower than the positivity threshold (titres in indirect fluorescent antibody test, IFAT, 80 to 160), which could be due to (1) infection is in the pre-patent period, (2) infection is in the remission period or (3) cross-reaction (Fisa et al. 1992).
For the diagnosis of HCL, direct detection of the parasite in skin samples is required (Giemsa's staining and/or cultures). The two main drawbacks of these techniques are the low number of parasites usually present in these samples and the invasive nature of a sampling technique such as biopsy. In immunocompromised subjects the humoral response to Leishmania infection is often negative (42.2%) (WHO, 1996) which limits the diagnostic value of serological techniques. Parasitological diagnosis in bone-marrow aspirates is the most commonly used technique and is among the most sensitive techniques (98%) compared to blood, normal skin, gastrointestinal tract, liver, spleen, pleural liquid and lymph nodes (WHO, 1996). However, this requires an invasive sampling method which is not advisable in individuals generally debilitated by HIV infection.
PCR-based methods for detecting Leishmania species in clinical samples have been developed which amplify rRNA and miniexon genes, kinetoplast DNA (kDNA) and repeated nuclear DNA sequences (Quiao, Miles & Wilson, 1995; Delgado et al. 1996; Ramos et al. 1996; Noyes et al. 1998). These methods are of variable specificity; some can detect all Leishmania species while other methods identify the infecting Leishmania parasite to the species level. These techniques mostly have a high sensitivity although some, such as PCR with a subsequent Southern blot hybridization (Uliana et al. 1994; Laskay et al. 1995; Andresen et al. 1996), are laborious and time consuming, and in practice are not very useful as routine diagnostic techniques. On the other hand, the diagnostic value of some of these PCR-based methods has not been confirmed on a sufficient number of parasite strains and clinical samples (Piarroux et al. 1993, 1995; Quiao et al. 1995).
In the present paper we describe a PCR–ELISA technique which can overcome the problems of the traditional diagnostic techniques, i.e. to (1) diagnose cases of HCL using less invasive sampling techniques than biopsy, (2) diagnose cases of HVL in HIV(+) individuals using peripheral blood samples instead of bone-marrow and (3) correctly meaning anti-Leishmania titres lower than the positivity threshold of CaL in dogs.
MATERIALS AND METHODS
L. infantum kDNA target and development of primers
A 196 bp EcoRI/HaeIII fragment diagnostic for L. infantum derived from a minicircle region that exhibits interspecies variation was used (Gramiccia et al. 1992, GenBank accession no. S49390). Sequence data were analysed and sets of primers for PCR were designed using the GCG Wisconsin Package software. Primers were synthesized by Pharmacia Biotech.
Strains and DNA preparation
A total of 33 Spanish strains of L. infantum from 19 different zymodemes that had been previously characterized by isoenzyme electrophoresis was used (Martín-Sánchez et al. 1996, 1999) (Table 1). Other reference strains used belong to the species L. donovani (1 strain), L. major (1), L. tropica (1), L. mexicana (1) and Trypanosoma cruzi (2).
Promastigotes were maintained in EMTM (WHO, 1989) and the parasites were grown to stationary phase. Genomic DNA was prepared by proteinase K digestion, phenol/chloroform extraction and ethanol precipitation (Sambrook, Fritsch & Maniatis, 1989). Human, dog and sandfly (Phlebotomus perniciosus) DNA were used as negative controls in the PCR assay.
PCR amplification
Primers 9 (forward primer: 5′-CAAAAGTCCCCACCAATCCC-3′) and 83 (reverse primer: 5′-AAACCCTGGTCTGGAGGCTTAG-3′) were used to amplify a 75 bp fragment contained within the 196 bp fragment. Reactions were carried out in 50 mM KCl, 10 mM Tris–HCl, pH 8.3, 1.5 mM MgCl2, 200 μM dNTPs, 12.5 pM each primer, 5–10 ng DNA or 0.5 μl samples, and 0.6 U Taq polymerase in a final volume of 25 μl. Each reaction was first denatured at 94 °C for 3 min and then cycled 40 times in a Progene Techne Thermal Cycler. Cycles were: 94 °C, 30 sec; 60 °C, 30 sec; 72 °C, 30 sec, with a final extension at 72 °C during 2 min. Amplified products were analysed by electrophoresis in 3.0% agarose gels, stained with ethidium bromide, and visualized under ultraviolet light.
Southern hybridization and DNA sequencing
Transfer of DNA from agarose gels to nylon membranes was performed as described by Sambrook et al. (1989). Labelling of DNA with digoxigenin hybridizations were carried out as described by Martín-Sánchez et al. (1998) using genomic DNA of one of the L. infantum strains (Table 1) and the amplified fragment of 121 bp as probes.
PCR products were purified by agarose gel fractionation and binding to glass milk. Sequencing was done using each of the PCR primers and the ABI PRISMTM BigDye Cycle Sequencing Ready Reaction kit.
PCR–ELISA
The amplification reaction was carried out using identical conditions to those used for PCR amplification except for the use of 2.5 μl of PCR labelling mix (Boehringer Mannheim) instead of the dNTP mixture. For ELISA detection of PCR products, we used the PCR–ELISA kit (DIG Detection) of Boehringer Mannheim, following the manufacturer's instructions. The oligonucleotide probe was selected from the 75 bp fragment without the primers necessary for its amplification. The oligonucleotide 5′-CCA AAC AGG GCA AAA ACC-3′ of Tm = 53.7 and %G+C = 50 was synthesized, purified and modified in position 5′ with biotin by Roche Diagnostics, LTD. Hybridization is carried out at 50 °C for 3 h agitation and washings carried out 3 times of 5 min duration each time. After incubation with the substrate (ABTSR), 40 min at room temperature with gentle shaking, absorbance values of plates were read at 405 and 492 nm (patent request number: P9901804).
Clinical samples and DNA preparation
Human samples were obtained from the following cases. (1) Three leishmaniasis asymptomatic patients coinfected with Leishmania-HIV. The presence of Leishmania in the bone-marrow was confirmed by optical microscopy of the Giemsa's-stained smears. PCR–ELISA was performed on samples of peripheral blood. (2) Seven cases of suspected HCL, 3 children under 6 years old and 4 adults. Skin samples were obtained by biopsy in 4 of these cases, in the other 3 cases they were obtained by scraping the edges of the lesion. (3) Two healthy adults from whom peripheral blood samples were used.
A total of 70 canine samples were obtained from 31 dogs who had been previously destined for euthanasia by the Society for the Protection of Animals and Plants of ‘Francisco de Asis' in Granada: 21 bone-marrow samples, 28 samples of peripheral blood, 18 samples of popliteal lymph node aspirate and 3 skin samples (biopsies of lesion margin).
DNA was prepared according to the technique described by D'Oliveira et al. (1995) followed by phenol/chloroform extraction and ethanol precipitation. With peripheral blood samples, buffy coat was carried out on an initial amount of 1–1.5 ml and the resulting product was divided into 2 equal parts for culture and DNA preparation.
Diagnostic techniques used as a reference
Simultaneously to and independent of PCR–ELISA, the following techniques for diagnosis of leishmaniasis were applied to the different clinical samples. (1) Human samples, (a) peripheral blood: culture; (b) skin, optical microscopy of samples stained with Geimsa's stain and cultured. (2) Canine samples, (a) bone-marrow, lymph node and skin: culture; (b) peripheral blood, culture and IFAT.
Parasite cultures were grown in EMTM medium (WHO, 1989) and kept for over 1 month before negative results were recorded. Positive cultures were mass cultured and the isolates were characterized by isoenzyme electrophoresis (Martín-Sánchez et al. 1994). The IFAT technique was carried out according to the description by Acedo et al. (1996).
RESULTS
Analysis of PCR amplification products
Amplification of 5–10 ng of genomic DNA of 3 strains of L. infantum (DP74, DP152 and DP204) generated 2 main products of 75 and 121 bp (Fig. 1) determined by DNA sequencing. The 75 bp fragment had 98.7% identity with the fragment theoretically designed using the probe of Gramiccia et al. (1992) (only one of 75 bp was different). The sequence of the 75 bp fragment was contained within the 121 bp fragment (Fig. 2). The amplification of this band of higher size is probably generated by the presence in the L. infantum kDNA minicircle, upstream of the primer 9 annealing site, of a region with high homology to the 3′ end of the primer 9. Since the annealing temperature used in PCR experiment was higher than Tm of the primer 9 we suggest that at least a large primer proportion is present upstream of the 75 bp minicircle fragment. These same bands were amplified when the remaining 30 strains of L. infantum (Table 1) were used. For some strains, other bands corresponding to fragments larger than 121 bp were amplified. All the amplification products of the 33 strains of L. infantum hybridized with the 2 probes used (genomic DNA and the 121 bp fragment) (data not shown). Amplification of 5–50 ng of genomic DNA of the other Leishmania species, including L. donovani, and T. cruzi, did not generate any product. Amplification was also negative when we used host DNA (man, dog, sandfly) free from L. infantum DNA (data not shown).
Fig. 1. PCR amplification products generated using genomic DNA of 3 strains of Leishmania infantum. Lane 1, PCR negative control; Lane 2, DP74 strain; Lane 3, DP152 strain; Lane 4, DP204 strain; Lane 5, DNA molecular weight marker V of Boehringer Mannheim.
Fig. 2. Alignment of the sequences of 75 bp, 121 bp and the 196 bp fragment of Gramiccia et al. (1992). (EMBL Nucleotide Sequence Database Accession Number: AJ272098 LIN272098 is not confidential). The primer 9 nucleotide sequence located in the 5′ end region of the 121 bp fragment is shown in bold.
Sensitivity of the PCR assay
Serial dilutions of 10 ng L. infantum genomic DNA were subjected to PCR and the PCR products were analysed in agarose gel containing ethidium bromide. The assay was positive, with amplification of the 75 and 121 bp fragments up to dilution 1/103, which is equivalent to the detection of approximately 10 pg of total DNA. Lower amounts of input DNA gave negative results. With Southern hybridization it was possible to detect 1 fg of genomic DNA (dilution 1/107) (Fig. 3). Also, raw extracts prepared from suspensions of promastigotes in culture were used for PCR, and the detection limit was 50–100 promastigotes (Fig. 4).
Fig. 3. Southern hybridization with the 121 bp DNA fragment as probe. Lane 1, PCR negative control; Lanes 2–9, Leishmania infantum DNA (10−7 to 10 ng).
Fig. 4. PCR amplification products generated using raw extracts prepared from suspensions of promastigotes in culture. Lane 1, PCR negative control; Lane 2, 1000 promastigotes; Lane 3, 500 promastigotes; Lane 4, 100 promastigotes.
ELISA detection of PCR products
This was designed to solve the two problems posed by detection by electrophoresis of PCR products i.e. (1) to increase the sensitivity and (2) amplification of fragments of higher molecular weight than the two main products of 75 and 121 bp.
The mean absorbance value (A405–A492) of the PCR control blank (PCR without DNA) was 0.04, and PCR with only host DNA (human or canine) was 0.1. The absorbance values obtained for L. donovani, L. tropica, L. mexicana and L. major were lower than 0.5 and for T. cruzi was below 0.2. For the 33 strains of L. infantum studied this was always greater than 3.0 (Fig. 5).
Fig. 5. Mean absorbance values (A405–A492) obtained with the PCR–ELISA; (a) 10 ng of genomic DNA of Leishmania infantum, (b) 0.01 ng of genomic DNA of L. infantum, (c) 1 fg of genomic DNA of L. infantum, (d) 0.1 fg of genomic DNA of L. infantum, (e) 1000 parasites of L. infantum, (f) 10 parasites of L. infantum, (g) 1 parasite of L. infantum, (h) 0.1 parasite of L. infantum, (i) mixtures with 0.1 parasite of L. infantum and host DNA (human and canine), (j) 0.01 parasite of L. infantum, (k) 10 ng of genomic DNA of L. donovani, (l) 10 ng of genomic DNA of L. major, (m) 10 ng of genomic DNA of L. tropica, (n) 10 ng of genomic DNA of L. mexicana, (o) 10 ng of genomic of DNA of Trypanosoma cruzi, (p) human DNA, (q) canine DNA, (r) phlebotomine DNA, (s) PCR negative control.
The detection limit of the PCR–ELISA assay performed with serial dilutions of 10 ng of L. infantum genomic DNA (DP74 strain) was 1 fg (dilution 1/107), and when this was determined using raw promastigote extract it corresponded to 0.1 promastigotes. In all cases, the absorbance readings (A405–A492) were always greater than 1.0. This corresponds to a rise in the sensitivity of PCR–ELISA in comparison with PCR detection of the amplified product by electrophoresis of 100–1000 times that obtained with PCR and ethidium bromide. Amplification of the mixtures with different proportions of raw extract of 1 promastigote, or L. infantum genomic DNA, and host DNA (human or canine) did not bring about changes in the absorbance levels compared to when L. infantum DNA was amplified in the absence of host DNA (Fig. 5: h and i).
Clinical samples: comparison of PCR–ELISA with other diagnostic techniques applied to HCL, HVL and CaL
The results obtained with the different samples of blood, bone-marrow, lymph node material and human or canine skin, with each of the diagnostic techniques used, are compiled in Tables 2 and 3. The reading was considered as positive when absorbance (A405–A492) was [ges ]1.
Table 2. Results obtained in the human clinical samples with each of the diagnostic techniques used (O.M., optical microscopy of the Giemsa's stained smears; P, positive result; N, negative result; p. blood, peripheral blood; b. marrow, bone marrow.)
Table 3. Results obtained in the canine clinical samples with each of the diagnostic technique used (1) Dog with cutaneous lesions: skin material obtained by biopsy was positive with PCR–ELISA and culture. N.P., Not performed; P, positive result, N, negative result.)
Human clinical samples (Table 2). The 7 patients with a clinical picture suggestive of HCL were positive with PCR–ELISA. Patient no. 1 had been diagnosed with HCL several years previously and had received specific treatment for this consisting of infiltration into the lesion of Glucantime and systemic administration of Allopurinol, without achieving complete remission. The culture was negative with 6 skin samples and only positive with the skin sample obtained from patient no. 7. However, optical microscopy of Giemsa's-stained smears gave positive results in patients nos. 4 and 7 on whom a biopsy had been performed, and negative in the remaining patients.
The 3 samples of peripheral blood from the 3 cases of HIV-Leishmania co-infection and with a positive bone-marrow Giemsa's-stained smear using optical microscopy, were positive with PCR–ELISA and negative in culture and Giemsa's-stained smear. The 2 healthy adults were negative with PCR–ELISA and also with the other diagnostic techniques uses.
Canine clinical samples (Table 3). Sixteen dogs (3 with IFAT 0 and 13 with IFAT [ges ]80), were confirmed as negative (the 3 former ones) or positive for all the diagnostic techniques performed (IFAT, culture and PCR–ELISA). Four dogs were negative in culture and positive with IFAT and PCR–ELISA. The 8 dogs with IFAT = 40 and 3 with IFAT = 20 were positive with PCR–ELISA and negative in culture. There was 100% agreement between the results obtained by PCR–ELISA using DNA from blood samples and DNA from lymph node material, and 88.2% between those obtained from blood and bone-marrow samples. The 3 skin samples analysed were positive in culture and with PCR–ELISA; the serology of the 3 animals was positive for CaL.
The parasites isolated from 10 of the 13 dogs with a positive culture were characterized by starch gel isoenzyme electrophoresis, and it was confirmed that they belonged to the species L. infantum and, more specifically, to 3 different zymodemes: GR-1 (MON-1), GR-11 (MON-199 var NP1) and GR-12 (MON-199) (Martín-Sánchez et al. 1999).
DISCUSSION
The PCR method described is based on the partial amplification of a cloned 196 bp fragment of L. infantum kDNA. When used as a probe in hybridization experiments with isotopic and non-isotopic systems this 196 bp fragment has been shown to be specific to L. infantum and to recognize the wide range of zymodemes of this species occurring around the Mediterranean basin (Gramiccia et al. 1992; Martín-Sánchez et al. 1998). The PCR was tested on 33 L. infantum strains belonging to 19 different zymodemes, and the amplified product was detected using 2 techniques, electrophoresis with ethidium bromide and ELISA, and the same results was obtained among strains. It is, therefore, very unlikely that false negatives would be produced because the target DNA would not be recognized in all L. infantum strains, as suggested by other authors. False negative PCR results were investigated by Piarroux et al. (1995) and it was demonstrated that the two main factors leading to such errors were the presence of inhibitory factors remaining in some samples after DNA preparation, and intra-specific polymorphism of the primer target DNA.
Sensitivity of the PCR–ELISA of L. infantum was 1 fg of genomic DNA. Because of this high sensitivity, this technique could be used to detect the presence of the parasite in samples of peripheral blood from dogs with CaL. L. infantum has rarely been detected in the circulating blood of the host (more specifically in the buffy coat), possibly because it is hardly ever practised with the exception of HIV(+) individuals who present a positivity of around 74.8% (WHO, 1996), Using culture techniques, Mathis & Deplazes (1995) detected Leishmania in leuco-concentrated blood of a dog with CaL but obtained negative results in another two. We also demonstrated the presence of Leishmania in leuco-concentrated blood from 2 dogs using cultures (Martín-Sánchez et al. 1998). However, we do not have data on the positivity rate in haemoculture.
Human clinical samples. The results suggest that it is possible to diagnose HCL by performing PCR–ELISA on samples obtained by scraping the lesion margins. Using this technique the problems of the current diagnostic methods for HCL can be avoided: the scar resulting from the biopsy and the low sensitivity of cultures and stained samples because of the low parasitic load. Indeed, culture, as a diagnostic technique, had a very low sensitivity in our study (1 out of 7), mainly due to the small number of parasites present but also because of the difficulty of growing some dermotropic strains in culture (Gramiccia & Gradoni, 1989; Gramiccia, Gradoni & Pozio, 1987). This also applies to the use of optical microscopy on the Giemsa's-stained samples: only 2 of the 7 patients gave positive results.
Interestingly, with regard to patients with HIV–Leishmania coinfection and haemoculture negative, the sensitivity of PCR–ELISA with peripheral blood samples was the same as that achieved with optical microscopy of the bone-marrow Giemsa's-stained smears. The results are interesting due to the use in this case of a non-invasive method. López-Vélez et al. (1995), in a study on 25 HIV–Leishmania coinfected patients with clinical symptoms, found a 67% positivity in haemocultures. It is possible that the difference in the results obtained is because these authors used 10 ml of blood in comparison to the 1–1.5 ml that we used for both techniques (culture and DNA preparation for PCR–ELISA), a volume that could be insufficient to obtain a positive result in culture. On the other hand, all 3 individuals included in our study were asymptomatic for VL. A larger number of samples is required in order to confirm these results.
Canine clinical samples. IFAT is the technique usually used for CaL diagnosis, and it is considered to be the reference method by the WHO. In this technique, threshold titres of 80 or 160 determine the presence or absence of the disease (Abranches et al. 1983; Bettini & Gradoni, 1986; Acedo-Sánchez et al. 1996; Morillas et al. 1996). All the samples we studied with titres equal or higher than these values gave positive results with PCR–ELISA. However, in our opinion the main problem concerns sera with titres below the threshold values (e.g. of 20 and 40). On several occasions, cultures have been used to isolate the parasite from dogs with titres of around 40 but in many others this has not occurred and cultures have proven to be of limited diagnostic value in these cases (Lanotte et al. 1978; Martínez-Cruz et al. 1990; Acedo-Sánchez et al. 1996). Moreover, cultures are often carried out on lymph node or bone-marrow samples, difficult to obtain, and involve a more traumatic invasion of host organs than simple extraction of blood samples. This is complicated further by the fact that cultures must be kept for at least 1 month to make a definitive negative diagnosis and this is both time consuming and costly.
Concordance between results by using PCR–ELISA on blood samples and those obtained when applying this same technique to other kinds of samples (bone-marrow and/or lymph node material) suggests the possibility of using the former for diagnosis of CaL, as occurs currently with HIV(+) individuals. Finally, it is noteworthy that the 3 dogs with titres of 20 were also positive with PCR–ELISA, whereas all those studied with a zero titre were negative with this technique.
We are grateful to the Subdirección General de Formación y Promoción del Conocimiento (PM97-0176) for financial support.
References
ABRANCHES, P., LOPES, F. J., SILVA, F. M. C., RIBIERO, M. M. S. & PIRES, C. A. (1983). Le kala-azar au Portugal. III. Résultats d'une enquête sur la leishmaniose canine réalisée dans les environs de Lisbonne. Comparison des zones urbaines et rurales. Annales Parasitologie Humaine et Comparée 58, 307–315.CrossRefGoogle Scholar
ACEDO-SANCHEZ, C., MARTIN-SANCHEZ, J., VELEZ-BERNAL, I. D., SANCHIS-MARIN, M. C., LOUASSINI, M., MALDONADO, J. A. & MORILLAS-MARQUEZ, F. (1996). Leishmaniasis eco-epidemiology in the Alpujarra region (Granada province, southern Spain). International Journal for Parasitology 25, 303–310.CrossRefGoogle Scholar
ANDRESEN, K., GAAFAR, A., EL-HASSAN, A. M., ISMAIL, A., DAFALLA, M., THEANDER, T. G. & KHARAZMI, A. (1996). Evaluation of the polymerase chain reaction in the diagnosis of cutaneous leishmaniasis due to Leishmania major: a comparison with direct microscopy of smears and sections from lesions. Transactions of the Royal Society of Tropical Medicine and Hygiene 90, 133–135.CrossRefGoogle Scholar
BETTINI, S. & GRADONI, L. (1986). Canine leishmaniasis in the Mediterranean area and its implications for human leishmaniasis. Insect Science and its Applications 7, 241–245.CrossRefGoogle Scholar
DELGADO, O., GUEVARA, P., SILVA, S., BELFORT, E. & RAMIREZ, J. L. (1996). Follow-up of a human accidental infection by Leishmania (Viannia) braziliensis using conventional immunological techniques and polymerase chain-reaction. American Journal of Tropical Medicine and Hygiene 55, 267–272.CrossRefGoogle Scholar
D'OLIVEIRA, C., VAN DER WEIDE, M., HABELA, M. A., JACQUIET, P. & JONGEJAN, F. (1995). Detection of Theileria annulata in blood samples of carrier cattle by PCR. Journal of Clinical Microbiology 33, 2665–2669.Google Scholar
FISA, R., PORTUS, M., GALLEGO, M., VALLAS, D. & AISA, M. J. (1992). El diagnóstico serológico de la leishmaniosis canina en la comarca del Priorat (Tarragona). Clínica Veterinaria de pequeños animales 12, 33–38.Google Scholar
GRAMICCIA, M. & GRADONI, L. (1989). Successful in vitro isolation and cultivation of italian dermotropic strains of Leishmania infantum sensu lato. Transactions of the Royal Society of Tropical Medicine and Hygiene 83, 76.CrossRefGoogle Scholar
GRAMICCIA, M., GRADONI, L. & POZIO, E. (1987). Leishmania infantum sensu lato as an agent of cutaneous leishmaniasis in Abruzzi region (Italy). Transactions of the Royal Society of Tropical Medicine and Hygiene 81, 235–237.CrossRefGoogle Scholar
GRAMICCIA, M., SMITH, D. F., ANGELICI, M. C., READY, P. D. & GRADONI, L. (1992). A kinetoplast DNA probe diagnostic for Leishmania infantum. Parasitology 105, 29–34.CrossRefGoogle Scholar
JAMBOU, D., MARTY, P., JAMBOU, R., LE FICHOUX, Y., HAAS, P., JOURDAIN, N. & BAYADA, M. (1986). Preliminary serological study on canine leishmaniasis in the Alpes Maritimes Department, France. Transactions of the Royal Society of Tropical Medicine and Hygiene 80, 666–667.CrossRefGoogle Scholar
LANOTTE, G., RIOUX, J. A., CROSET, H. & VOLHARD, Y. (1978). Ecologie des Leishmanioses dans le Sud de la France. Les méthodes d'echantillonnage dans le dépistage et l'analyse de l'enzootie canine. Annales de Parasitologie Humaine et Comparée 53, 33–45.CrossRefGoogle Scholar
LASKAY, T., MIKO, T. L., NEGRESSE, Y., SOLBACH, W., ROLLINGHOFF, M. & FROMMEL, D. (1995). Detection of cutaneous Leishmania infection in paraffin-embedded skin biopsies using the polymerase chain reaction. Transactions of the Royal Society of Tropical Medicine and Hygiene 89, 273–275.CrossRefGoogle Scholar
LOPEZ-VELEZ, R., LAGUNA, F., ALVAR, J., PEREZ-MOLINA, J. A., MOLINA, R., MARTINEZ, P. & VILLARRUBIA, J. (1995). Parasitic culture of buffy coat for diagnosis of visceral leishmaniasis in human immodeficiency virus-infected patients. Journal of Clinical Microbiology 33, 937–939.Google Scholar
MARTIN-SANCHEZ, J., GRAMICCIA, M., ANGELICI, M. C., ACEDO-SANCHEZ, C. & MORILLAS-MARQUEZ, F. (1998). Evaluation of the PLiD2-196bp kDNA probe specific for Leishmania infantum for the diagnosis of canine leishmaniasis in a focus of southern Spain. Research and Reviews in Parasitology 58, 91–94.Google Scholar
MARTIN-SANCHEZ, J., LEPE, J. A., TOLEDO, A., UBEDA, J. M., GUEVARA, D. C., MORILLAS-MARQUEZ, F. & GRAMICCIA, M. (1999). Leishmania infantum enzymatic variants causing canine leishmaniasis in the Huelva province (south-west Spain). Transactions of the Royal Society of Tropical Medicine and Hygiene 93, 495–496.CrossRefGoogle Scholar
MARTIN-SANCHEZ, J., MORILLAS-MARQUEZ, F., SANCHIZ-MARIN, M. C. & ACEDO-SANCHEZ, C. (1994). Isoenzymatic characterization of the etiologic agent of canine leishmaniasis in the Granada region of southern Spain. American Journal of Tropical Medicine and Hygiene 50, 758–762.CrossRefGoogle Scholar
MARTIN-SANCHEZ, J., RUIZ-MARTINEZ, F., SALINAS-MARTINEZ DE LECEA, J. M., SANCHEZ-RABASCO, C., ACEDO-SANCHEZ, C., SANCHIZ-MARIN, M. C., DELGADO-FLORENCIO, V. & MORILLAS-MARQUEZ, F. (1996). Leishmania infantum Nicolle, 1908 from southern Spain: Characterisation of the strains from human visceral and cutaneous leishmaniasis and from sandflies; with a numerical analysis of the isoenzymatic data. Systematic Parasitology 33, 177–182.CrossRefGoogle Scholar
MARTINEZ-CRUZ, S., MARTINEZ-MORENO, A., MARTINEZ-MORENO, F. J., MARTINEZ-GÓMEZ, F. & HERNANDEZ-RODRIGUEZ, S. (1990). Epidemiología de la Leishmaniosis canina en Córdoba. Revista Ibérica de Parasitologia 50, 1–7.Google Scholar
MATHIS, A. & DEPLAZES, P. (1995). PCR and in vitro cultivation for detection of Leishmania spp. In diagnostic samples from humans and dogs. Journal of Clinical Microbiology 33, 1145–1149.Google Scholar
MORILLAS, F., SANCHEZ-RABASCO, F., OCAÑA, J., MARTIN-SANCHEZ, J., OCAÑA-WIHELMI, J., ACEDO, C. & SANCHIZ-MARIN, M. C. (1996). Leishmaniosis in the focus of the Axarquía region, Malaga province, southern Spain: a survey of the human, dog, and vector. Parasitology Research 82, 569–570.CrossRefGoogle Scholar
NIETO, C. G., NAVARRETE, M., HABELA, M. & HERNANDEZ-RODRIGUEZ, S. (1992). Seroprevalence of canine leishmaniasis around Cáceres, Spain. Preventive Veterinary Medicine 13, 173–178.CrossRefGoogle Scholar
NOYES, H. A., REYBURN, H., BAILEY, J. W. & SMITH, D. (1998). A nested-PCR-based schizodeme method for identifying Leishmania kinetoplast minicircle classes directly from clinical samples and its application to the study of the epidemiology of Leishmania tropica in Pakistan. Journal of Clinical Microbiology 36, 2877–2881.Google Scholar
PIARROUX, R., AZAIEZ, R., LOSSI, A. M., REYNIER, P., MUSCATELLI, F., GAMBARELLI, F., FONTES, M., DUMON, H. & QUILICI, M. (1993). Isolation and characterization of a repetitive DNA sequence from Leishmania infantum: development of a visceral leishmaniasis polymerase chain reaction. American Journal of Tropical Medicine and Hygiene 49, 364–369.CrossRefGoogle Scholar
PIARROUX, R., FONTES, M., PERASSO, R., GAMBARELLI, F., JOBLET, C., DUMON, H. & QUILICI, M. (1995). Phylogenetic relationships between Old World Leishmania strains revealed by analysis of a repetitive DNA sequence. Molecular and Biochemical Parasitology 32, 746–749.CrossRefGoogle Scholar
QUIAO, Z., MILES, M. A. & WILSON, S. M. (1995). Detection of parasites of the Leishmania donovani complex by a polymerase chain reaction-solution hybridization enzyme-linked immunoassay (PCR–SHELA). Parasitology 110, 269–275.CrossRefGoogle Scholar
RAMOS, A., MASLOV, D. A., FERNANDES, O., CAMPBELL, D. A. & SIMPSON, L. (1996). Detection and identification of human pathogenic Leishmania and Trypanosoma species by hybridization of PCR-amplified mini-exon repeats. Experimental Parasitology 82, 242–250.CrossRefGoogle Scholar
ULIANA, S. R. B., NELSON, K., BEVERLEY, S. M., CAMARGO, E. P. & LOETER-WINTER, L. M. (1994). Discrimination amongst Leishmania by polymerase chain reaction and hybridization with small subunit ribosomal DNA derived oligonucleotides. Journal of Eukaryotic Microbiology 41, 324–330.CrossRefGoogle Scholar
SAMBROOK, J., FRITSCH, E. F. & MANIATIS, T. (1989). Molecular Cloning. A Laboratory Manual, 2nd Edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, USA.
SEMIAO-SANTOS, S. J., EL HARITH, A., FERREIRA, E., PIRES, C. A., SOUSA, C. & GUSMAO, R. (1995). Evora district as a new focus for canine leishmaniasis in Portugal. Parasitology Research 81, 235–239.Google Scholar
WORLD HEALTH ORGANIZATION (1989). Handbook on isolation, characterization and cryopreservation of Leishmania. UNDP/World Bank/WHO Special Programme for Research and Training in Tropical Diseases. Ed. David Evans. WHO, Geneva.
WORLD HEALTH ORGANIZATION (1996). Epidemiological analysis of 692 retrospective cases of Leishmania/HIV co-infection. World Health Organization. Division of Control of Tropical Diseases. WHO/LEISH/96.39. WHO, Geneva.
Table 1. Leishmania infantum isolates used in this study and zymodeme data
Fig. 1. PCR amplification products generated using genomic DNA of 3 strains of Leishmania infantum. Lane 1, PCR negative control; Lane 2, DP74 strain; Lane 3, DP152 strain; Lane 4, DP204 strain; Lane 5, DNA molecular weight marker V of Boehringer Mannheim.
Fig. 2. Alignment of the sequences of 75 bp, 121 bp and the 196 bp fragment of Gramiccia et al. (1992). (EMBL Nucleotide Sequence Database Accession Number: AJ272098 LIN272098 is not confidential). The primer 9 nucleotide sequence located in the 5′ end region of the 121 bp fragment is shown in bold.
Fig. 3. Southern hybridization with the 121 bp DNA fragment as probe. Lane 1, PCR negative control; Lanes 2–9, Leishmania infantum DNA (10−7 to 10 ng).
Fig. 4. PCR amplification products generated using raw extracts prepared from suspensions of promastigotes in culture. Lane 1, PCR negative control; Lane 2, 1000 promastigotes; Lane 3, 500 promastigotes; Lane 4, 100 promastigotes.
Fig. 5. Mean absorbance values (A405–A492) obtained with the PCR–ELISA; (a) 10 ng of genomic DNA of Leishmania infantum, (b) 0.01 ng of genomic DNA of L. infantum, (c) 1 fg of genomic DNA of L. infantum, (d) 0.1 fg of genomic DNA of L. infantum, (e) 1000 parasites of L. infantum, (f) 10 parasites of L. infantum, (g) 1 parasite of L. infantum, (h) 0.1 parasite of L. infantum, (i) mixtures with 0.1 parasite of L. infantum and host DNA (human and canine), (j) 0.01 parasite of L. infantum, (k) 10 ng of genomic DNA of L. donovani, (l) 10 ng of genomic DNA of L. major, (m) 10 ng of genomic DNA of L. tropica, (n) 10 ng of genomic DNA of L. mexicana, (o) 10 ng of genomic of DNA of Trypanosoma cruzi, (p) human DNA, (q) canine DNA, (r) phlebotomine DNA, (s) PCR negative control.
Table 2. Results obtained in the human clinical samples with each of the diagnostic techniques used
Table 3. Results obtained in the canine clinical samples with each of the diagnostic technique used
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