Evidence for Leishmania (Viannia) Parasites in the Skin and Blood of Patients Before and After Treatment

Transcription

1 MAJOR ARTICLE Evidence for Leishmania (Viannia) Parasites in the Skin and Blood of Patients Before and After Treatment Carolina Vergel, Ricardo Palacios, a Horacio Cadena, Claudia Jimena Posso, Liliana Valderrama, Mauricio Perez, John Walker, Bruno Luis Travi, a and Nancy Gore Saravia Centro Internacional de Entrenamiento e Investigaciones Medicas, Cali, Colombia Background. American cutaneous leishmaniasis is considered to be a zoonotic disease transmitted by sand flies that feed on infected sylvatic mammals. However, the domestication of transmission and the increase in treatment failure with antimonial drugs have raised the suspicion of anthroponotic transmission of American cutaneous leishmaniasis. Methods. The objective of the present study was to explore the potential of humans as a source of infection for sand flies. Biological (xenodiagnosis and culture) and molecular (polymerase chain reaction/southern blot) detection methods were used to evaluate peripheral-blood monocytes and tissue fluids from sites accessible to sand flies from 59 adult patients with parasitologically confirmed American cutaneous leishmaniasis. Results. Overall, 44.1% of patients (26/59) presented biological and/or molecular evidence of Leishmania parasites in normal skin, peripheral-blood monocytes, lesion scars, or lesion border (by xenodiagnosis) before (18/ 59 [30.5%]) or after (10/27 [37.0%]) treatment. Leishmania parasites were cultured from the unaffected skin of 2 (3.6%) of 55 patients, and xenodiagnosis gave positive results for 5 (8.8%) of 57 patients before treatment. Conclusions. The presence of Leishmania parasites in the unaffected skin and peripheral-blood monocytes of a high proportion of patients even after treatment and the acquisition of infection by sand flies support the plausibility of anthroponotic transmission of American cutaneous leishmaniasis. American cutaneous leishmaniasis has long been considered a zoonosis, with humans being accidental hosts and a dead end in the cycle of transmission. Indisputably, all New World Leishmania parasites have a natural enzoonotic cycle of transmission. However, changing demographic patterns, including the massive settlement of tropical rain forests, have perturbed the Received 13 January 2006; accepted 20 March 2006; electronically published 10 July Presented in part: 53rd Congress of American Society of Tropical Medicine and Hygiene (poster 687), Miami, Florida, 7 11 November Potential conflicts of interest: none reported. Financial support: Burroughs Wellcome Wellcome Trust Infectious Disease Initiative, United Kingdom (grant /Z/99/Z); COLCIENCIAS, Colombia (grant ). R.P. is supported by a studentship grant from the Brazilian National Council for Scientific and Technological Development. a Present affiliations: Division of Infectious Diseases, Federal University of São Paulo UNIFESP, São Paulo, Brazil (R.P.); University of Texas Health Science Center, San Antonio (B.L.T.). Reprints or correspondence: Dr. Nancy G. Saravia, Centro Internacional de Entrenamiento e Investigaciones Medicas, Av. 1 Norte no. 3-03, Cali, Valle, Colombia (saravian@cideim.org.co). The Journal of Infectious Diseases 2006; 194: by the Infectious Diseases Society of America. All rights reserved /2006/ $15.00 natural cycle of transmission and led to intra- and peridomestic transmission and the urbanization of transmission in several countries of South America [1 5]. This trend has been marked by a shift in age-specific prevalence toward increases in those of younger ages, notably in children of preschool age and infants. Contemporaneously with this epidemiological change, the clinical treatment failure of antimonial (Sb) drugs has become increasingly evident [6 8]. Therapeutic response depends on several factors, including host immune response to infection [9, 10], pharmacokinetics, adherence to treatment [7], and drug quality, as well as the sensitivity of the parasite to Sb drugs [11, 12]. Nevertheless, the high frequency of treatment failure in Bihar, India [11], where transmission is anthroponotic, and its association with decreased in vitro sensitivity to Sb drugs [11] raise suspicion that resistant organisms may be propagated by humans who receive inadequate treatment and, consequently, harbor and transmit less susceptible strains of Leishmania to sand flies. Previous studies have shown that Lutzomyia longi Leishmania in Skin and Blood JID 2006:194 (15 August) 503

2 palpis, a surrogate vector of Leishmania (Viannia) parasites, could acquire infection when fed on the border of dermal lesions from patients with active disease [13, 14]. In addition, Leishmania kinetoplast DNA has been detected by polymerase chain reaction (PCR) in healed lesions several years after treatment and in peripheral blood in the absence of active disease [15, 16], and parasites have been isolated from healed lesions [17]. These findings and the demonstration of reactivation as a cause of recurrent episodes of cutaneous leishmaniasis [18] substantiate the persistence of L. (Viannia) parasites in previously affected skin sites and blood in the absence of disease. However, the epidemiological significance of persistent subclinical infection by dermotrophic Leishmania parasites is unknown. We conducted the present study to determine whether Leishmania parasites are present in sites and tissues of infected humans that are accessible to sand flies seeking a blood meal. Furthermore, we sought to determine whether Leishmania prevalence in these tissues would be altered by treatment. MATERIALS AND METHODS Study design. An exploratory study was conducted to establish thresholds of frequency, presumptively demarcating the biological relevance of Leishmania parasites in tissues accessible to sand flies. The sample size was calculated to be 49 subjects, with an expected prevalence of the phenomenon of 15%, a precision of 10%, and a significance level of 5%. This number was increased by 20% to 59 subjects, to account for potential losses due to incomplete or inadequate samples and patient attrition. Patients who received standard pentavalent Sb (Sb V ) treatment and participated in the pretreatment procedures to detect Leishmania parasites were invited to participate in the repeat of those procedures 13 weeks after the initiation of treatment. The study protocol and informed consent were approved by the Centro Internacional de Entrenamiento e Investigaciones Medicas (CIDEIM) Institutional Review Board for the protection of human subjects from research risks and were in accordance with the guidelines of the Declaration of Helsinski, good clinical practice, and the Colombian Ministry of Social Protection (Resolution 8430; 1993). Written, informed consent was obtained from all participants or their parents or guardians after the explanation of study procedures, potential benefits and risks, and the voluntary nature of participation. Inclusion and exclusion criteria. Patients who had parasitologically confirmed cutaneous leishmaniasis of 3 months of evolution and who were 16 years of age were eligible to participate. Patients were excluded from the study if they had previously received a diagnosis of leishmaniasis, had mucosal involvement, were pregnant or lactating, had received Sb V drugs for their current lesions, presented relative or absolute contraindication for the use of Sb V drugs (including a history of cardiac, renal, or hepatic disease), or had inappropriate localization of a cutaneous lesion or lymphatic region for direct xenodiagnosis (e.g., earlobe or eye). Patients. Fifty-nine patients with active cutaneous leishmaniasis of 3 months of evolution who consulted the CID- EIM diagnostic facilities in Cali or the Hospital San Andres in Tumaco were enrolled in the study. Of these, 27 were available 13 weeks after the initiation of treatment for clinical evaluation and repeated sampling, to detect persistent infection. Sb V treatment and follow-up. After diagnosis and the completion of initial study procedures, meglumine antimoniate (Glucantime; Aventis Farma) was administered intramuscularly at a dosage of 20 mg/kg/day for 20 days, in accordance with Pan American Health Organization/World Health Organization and Colombian Ministry of Social Protection guidelines for the treatment of cutaneous leishmaniasis. Glucantime was supplied by the Colombian Ministry of Social Protection. Initial therapeutic response was evaluated at 13 weeks after the initiation of treatment. Patients who experienced therapeutic failure defined as an increase in legion size, the appearance of new lesions, incomplete epithelialization, and/or persistent signs of inflammation before the 13th week were reevaluated clinically, and laboratory samples were obtained (by direct smear, culture of aspirate, or 4-mm-punch biopsy) to confirm the diagnosis and identify Leishmania species. Re-treatment with meglumine antimoniate was administered in accordance with national guidelines. Study procedures. The procedures for the detection of the presence of Leishmania parasites before and after treatment are schematically summarized in figure 1. Normal skin sites were defined as those proximal to a path of lymphatic drainage, at a distance of cm from a cutaneous lesion. Samples from scars were obtained after the completion of treatment ( 13 weeks after the initiation of therapy) and only from patients who were deemed to be clinically cured. Separation of monocytes. Monocytes were isolated and purified from 10 ml of human peripheral blood by use of Nyco- Prep (1.068 g/ml; Axis-Shield), in accordance with the manufacturer s instructions. After washing and centrifugation, the 6 6 pellet ( cells) was resuspended in 1000 ml of Dulbecco s PBS (DPBS; Gibco-BRL); 100 ml was cultured in duplicate in Senekjie s diphasic culture medium for isolation of parasites, 300 ml (containing ,000 monocytes) was used to artificially feed sand flies (indirect xenodiagnosis), and the rest (500 ml) was used to detect Leishmania parasites by molecular methods. For the latter, monocytes were centrifuged at 1300 g for 5 min, resuspended in 200 ml of DPBS, and frozen at 20 C until processed. Direct xenodiagnosis. Two body sites were concomitantly evaluated: the border of an active lesion and a normal skin site. 504 JID 2006:194 (15 August) Vergel et al.

3 Figure 1. Both the lesion and the normal skin site were cleaned with 70% alcohol. Four 50-mL polystyrene tubes (Falcon) that were covered by a fine mesh and that contained 25 female sand flies (Lu. longipalpis) each were placed against the selected site. Feeding was allowed ad libitum for 30 min. After feeding, the insects were maintained at 26 C and 85% 90% humidity, exclusively on a sucrose diet. Five to seven days later, the insects were dissected to search for Leishmania promastigotes in the gut [13, 14]. Indirect xenodiagnosis. Monocytes were used for artificial feeding with 100 female sand flies (Lu. longipalpis), as described by Ward et al. [19]. Approximately 300, ,000 cells were mixed with 2 ml of rabbit blood that had been inactivated at 56 C for 25 min and then placed in a vial covered with chickskin membrane; the flies were fed for 45 min. Under these experimental conditions, human monocytes would be ingested by a fully engorged sand fly ( 0.5 ml) during indirect xenodiagnosis, producing a concentration similar to that of monocytes in peripheral blood. Both methods of xenodiagnosis were performed before and after treatment. Culture of dermal tissue fluid aspirates and monocytes. Dermal tissue fluid for culture and PCR was obtained as described elsewhere [20]. Dermal tissue fluid aspirates and peripheral-blood monocytes were inoculated into Senekjie s diphasic culture medium [20]. PCR/Southern blot. Extraction of DNA and amplification of L. (Viannia) minicircle kinetoplast DNA (kdna) from dermal tissue fluid aspirates and peripheral-blood monocytes before and after treatment were conducted as described elsewhere [21]. PCR was performed with the primers B1 (5 -GGG GTT GGT GTA ATA TAG TGG-3 ) and B2 (5 -CTA ATT GTG CAC GGG GAG G-3 ), which have been previously reported to specifically amplify the entire 750-bp minicircle kdna of L. Schematic summary of the experimental strategy (Viannia) species [22, 23]. To improve sensitivity and specificity, a chemiluminescent Southern blot was done at high stringency (65 C) on hybridized PCR products as described elsewhere [21], using a probe comprising the 750-bp PCR product derived from L. panamensis kdna in combination with the Alka-Phos Direct labeling and detection system and CDP-Star (Amersham-Pharmacia Biotech), in accordance with the manufacturer s instructions. Identification of Leishmania species. Parasites isolated from skin lesions, normal skin, or scars were identified by indirect immunofluorescence with monoclonal antibodies [24]. Identification on the basis of isoenzyme polymorphisms was undertaken if strains could not be identified on the basis of reactivity with the panel of monoclonal antibodies [25]. Statistical analyses. Statistical analyses were performed using SPSS for Windows (version 7.5; SPSS). Associations between frequencies of positive samples by biological and molecular methods and demographic and clinical characteristics of the patients from whom the samples were isolated were determined by use of contingency tables and the x 2 test. Relationships between the number of evaluable sand flies after xenodiagnosis (surviving, engorged specimens) and the acquisition of infection were examined by use of the Kruskal-Wallis test followed by the Dunn procedure. P!.05 was considered to be statistically significant. RESULTS The 59 patients were predominantly males of African or mestizo ethnicity and were between 16 and 55 years of age (table 1). The majority resided in areas where leishmaniasis transmission was endemic. Regional adenopathy was detected in 45 (76.3%) of the patients at diagnosis (table 1). Twenty-seven patients Leishmania in Skin and Blood JID 2006:194 (15 August) 505

4 Table 1. Demographic and clinical characteristics of the study patients. Characteristic Before treatment (n p 59) After treatment (n p 27) Sex Male 54 (91.5) 26 (96.3) Female 5 (8.5) 1 (3.7) Race Black 30 (50.8) 13 (48.1) Mixed 21 (35.6) 11 (40.7) White 5 (8.5) 3 (11.1) Indigenous 2 (3.4) No data 1 (1.7) Adenopathy Yes 45 (76.3) 23 (85.2) No 10 (16.9) 4 (14.8) No data 4 (6.8) Reside in endemic area Yes 44 (74.6) 19 (70.4) No 14 (23.7) 8 (29.6) No data 1 (1.7) Age years 15 (25.4) 7 (25.9) years 15 (25.4) 5 (18.5) years 8 (13.6) 4 (14.8) years 12 (20.3) 5 (18.5) years 3 (5.1) 2 (7.4) 140 years 6 (10.2) 4 (14.8) Median (range), years 25.0 (16 55) 27 (17 55) Time of lesion evolution, median (range), months 1.5 ( ) Lesions per patient, median (range), no. 2.0 (1 15) NOTE. Data are no. (%) of patients, unless otherwise specified. (45.8%) provided samples at follow-up as well as before treatment; the demographic characteristics of these patients were similar to those of the patients who were unavailable for followup (table 1). No associations were found between evidence of the presence of Leishmania parasites by biological or molecular methods and the demographic or clinical characteristics of the patients. Leishmania parasites were isolated and identified at diagnosis from 40 (67.8%) of the 59 patients. Thirty-eight (95.0%) of the isolates were identified as L. panamensis, 1 (2.5%) was identified as L. braziliensis, and 1 (2.5%) was identified as L. (Viannia) species. In total, 26 (44.1% [95% confidence interval {CI}, 31.2% 57.6%]) of the 59 patients presented biological or molecular evidence of Leishmania parasites in tissues accessible to sand flies; 18 (30.5% [95% CI, 19.2% 43.9%]) of 59 patients presented evidence before treatment, and 10 (37.0% [95% CI, 19.4% 57.6%]) of 27 patients presented evidence after treatment. The frequency of evidence of Leishmania infection varied according to the method of detection (molecular or biological), sample type (dermal fluid or peripheral-blood monocytes), and dermal site (normal skin, active lesion, or scar) (table 2). PCR of peripheral-blood monocytes with confirmation by Southern blot was the most sensitive procedure, producing positive results in 25.9% of patients overall and in 19.2% (95% CI, 9.6% 32.5%) and 17.4% (95% CI, 5.0% 38.8%) of patients before treatment and at follow-up, respectively. Leishmania kdna was amplified from tissue fluid from normal skin in 7 (12.7%) of 55 patients (figure 2). The proportion of positive dermal tissue fluid aspirates from unaffected skin was higher after treatment (3/22; 13.6% [95% CI, 2.9% 34.9%]) than before treatment (4/53; 7.5% [95% CI, 2.1% 18.2%]) (table 2), but the difference was not significant. Notably, the rates of positive PCR/ Southern blot assays for normal skin (3/22; 13.6% [95% CI, 2.9% 34.9%]), peripheral-blood monocytes (4/23; 17.4% [95% CI, 5.0% 38.8%]), and scars (4/22; 18.2% [95% CI, 5.2% 40.3%]) were similar 13 weeks after the initiation of treatment (table 2). Direct biological evidence of the infectiousness of patient lesions before treatment was obtained for 5 (8.8% [95% CI, 2.9% 19.3%]) of 57 patients who transmitted parasites to sand flies that fed on the lesion border. Direct xenodiagnosis before treatment and indirect xenodiagnosis (by artificial feeding) at follow-up was negative for normal skin and healed lesions. The number of sand flies that fed during xenodiagnosis varied from 2to190. A significantly larger number of sand flies fed on lesion borders than on normal skin or healed lesions ( P!.05). Notably, xenodiagnosis was positive only when 50 sand flies had engorged ( P!.001) (table 3), and the proportion of positive results increased to 31.3% (5/16) when at least 50 sand flies had fed and survived for evaluation. In addition, Leishmania parasites were cultured from 2 (3.6% [95% CI, 0.4% 12.5%]) of 55 dermal tissue fluid aspirates obtained from unaffected skin during active disease. DISCUSSION The increasing importance of domestic transmission of cutaneous leishmaniasis [1, 26, 27] and clinical treatment failure due, at least in part, to drug-resistant parasites [12] support the plausibility of humans as reservoirs of Leishmania parasites. Evidence that humans can act as a reservoir of New World leishmaniases is limited to xenodiagnosis of patients and seropositive inhabitants of an area in Brazil where L. chagasi transmission is endemic [28] and to anecdotal reports of xenodiagnosis on the border of cutaneous lesions: in Venezuela, L. braziliensis using the natural vector Lu. youngi [29]; in Brazil, L. amazonensis using the surrogate vector Lu. longipalpis but 506 JID 2006:194 (15 August) Vergel et al.

5 Table 2. Methods of detection of Leishmania parasites and sources of the parasites in positive patients. Before treatment Patient Molecular methods (PCR/Southern blot) Normal skin Monocytes Biological methods Xenodiagnosis of lesion border Culture of normal skin After treatment, molecular methods (PCR/Southern blot) Normal skin Monocytes Scar NS NS NS 3543 NS NS NS NS NS NS NS NS NS 3847 NS NS + NS NS NS NS NS NS NS NS NS NS + NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS Subtotal by detection method/source 4/53 (7.5%) 10/52 (19.2%) 5/57 (8.8%) 2/55 (3.6%) 3/22 (13.6%) 4/23 (17.4%) 4/22 (18.2%) Subtotal by detection method a 13/57 (22.8%) 7/59 (11.9%) 7/27 (25.9%) Subtotal by before/after treatment 18/59 (30.5%) 10/27 (37.0%) Total 26/59 (44.1%) NOTE. NS, not sampled; PCR, polymerase chain reaction. a Excluding data on scars. not L. braziliensis using either Lu. longipalpis or its natural vector, Lu. intermedia [30]; and in Colombia, L. panamensis using Lu. longipalpis [14]. However, several studies have detected the presence of live L. (Viannia) parasites in blood [16] and in scars [17] even several years after healing, and Leishmania DNA has been amplified from peripheral-blood cells and from scars [15, 31, 32]. The present study was designed to demonstrate the availability of L. (Viannia) parasites in tissues accessible to sand flies in the human host. The results show that at least 44% of patients with cutaneous leishmaniasis present molecular and/ or parasitological evidence of Leishmania parasites in blood, normal skin, or healed lesions, in addition to the active lesion. Treatment did not diminish the overall rate of positivity. However, biological methods did not yield parasites in the 22 patients evaluated after treatment. The low sensitivity of the biological methods as used and the reduced number of patients available for evaluation after treatment do not allow conclusions to be drawn about the impact that treatment has on infectivity. Remarkably, Leishmania parasites were detected in normal skin before (7.5%) and after (13.6%) treatment by molecular methods and were also cultured from the normal skin of 2 patients before treatment, hence proving that viable parasites Leishmania in Skin and Blood JID 2006:194 (15 August) 507

6 Figure 2. Polymerase chain reaction amplification of Leishmania kinetoplast DNA (kdna) in dermal tissue fluid aspirated from the normal skin of patients with American cutaneous leishmaniasis, using L. (Viannia) specific primers B1 and B2. Lane 1, Positive control (10 pg of kdna); lane 2, positive control (1 pg of kdna); lane 3, patient 6579; lane 4, patient 3510; lane 5, patient 6600; lane 6, patient 6418; lane 7, negative control (ultrapure water); lane 8, molecular-weight marker (FX174); lane 9, patient 6459; lane 10, patient 6398; lane 11, patient 6552; lane 12, patient 3528; lane 13, patient 6573; lane 14, patient are present in normal skin, not only in the lesion. To our knowledge, this is the first demonstration of L. (Viannia) parasites in normal skin, supporting the notion that these parasites are available in the unaffected skin of individuals with active disease. Together with the frequent presence of Leishmania DNA in peripheral-blood monocytes before and after treatment, the demonstration of parasites in extralesional tissues shows that L. (Viannia) parasites circulate throughout the body during active disease and after the clinical resolution of lesions. Five patients (8.6%) had positive xenodiagnostic results before treatment. In light of the small number of sand flies that fed on the lesion borders and normal skin of individual patients during two 30-min intervals, compared with the permanent exposure to sand-fly bites in endemic foci of transmission, our results provide a minimal estimate of the infectiousness of patients for sand flies. Children with L. chagasi infection in Brazil have been found to be 11 times more infectious for sand flies than adults [28]. As in L. chagasi transmission, L. (Viannia) infection is more likely to cause disease in children than in adults that is, they are more susceptible [33]. In consequence, the exclusion of children from this study (because of concern about tolerability) is likely to have contributed to an underestimation of the ability of the affected human population to infect sand flies, particularly in circumstances of peri- and intradomiciliary transmission, where they constitute a large and growing proportion of patients. Xenodiagnostic experiments conducted in dogs have shown that individual hosts vary in infectiousness; a few (7/42) infected dogs proved to be highly infectious, accounting for 180% of the sand-fly infections, whereas others did not infect sand flies under the experimental conditions used [34]. PCR and culture of seropositive blood donors from an area in which L. infantum/chagasi transmission was endemic have demonstrated intermittent parasitemia in asymptomatically infected individuals [35 37]. Hence, although parasitemia may be more frequently demonstrable in symptomatic patients or dogs, asymptomatically infected individuals are also a plausible source of infection [9, 38]. The use of a species of sand fly that is a natural vector of the L. (Viannia) subgenus as well as engorgement of a larger number of sand flies might improve the sensitivity of xenodiagnosis. Xenodiagnosis of L. braziliensis was achieved before treatment in 8 patients by feeding Lu. youngi, a natural vector of this species in Venezuela, on the borders of lesions of 1 3 months duration [28]. An impressive 3.6% 19.2% of flies became infected [28]. In the present study, positive xenodiagnosis was only achieved when at least 50 sand flies had fed and survived until evaluation 5 days later. If we considered in the analysis only patients from whom 50 sand flies were available for evaluation after the flies had been fed on the lesion border, the proportion of positive xenodiagnostic results increased to 31.3% (5/16). Fifty or more sand flies were available for evaluation after xenodiagnosis on normal skin or after artificial feeding on monocytes for very few patients either before (6 and 4 patients, respectively) or after (0 and 3 patients, respectively) treatment (table 3). Therefore, the sensitivity of the xenodiagnostic procedure for normal skin and monocytes or for healed lesions was not comparable with that of the xenodiagnostic procedure for lesion borders, and, consequently, the results with regard to the infectivity of patients after treatment are inconclusive. The reason that sand flies fed more readily when exposed to lesion borders than to unaffected skin is unknown. However, because parasite burden and the density of inflammatory mononuclear cells harboring parasites are likely to be higher at the lesion border, a preference for feeding on or near lesions in nature would favor acquisition of infection. Molecular methods for the detection of genetic material have proven to be more sensitive than biological methods for the detection of Leishmania parasites themselves [39, 40]. Consequently, such methods are largely replacing less sensitive parasitological strategies for investigating the natural history and epidemiology of leishmaniasis and other transmissible diseases. In the present evaluation of the presence and availability of Leishmania parasites to sand-fly vectors, the specificity and enhanced sensitivity of PCR coupled with DNA hybridization demonstrated in a previous study [21] was verified the presence of Leishmania parasites was detected at a higher frequency (21/59 [35.6%]) by this method than by all biological methods combined (7/59 [11.9%]). Although the molecular-detection 508 JID 2006:194 (15 August) Vergel et al.

7 Table 3. No. of Lutzomya longipalpis sand flies used for direct and indirect xenodiagnosis of cutaneous leishmaniasis caused by Leishmania (Viannia) species. Xenodiagnostic method, source Patients evaluated, total no. Surviving fed sand flies Median (range), no. a Kruskal-Wallis P Total no. Patients with 50 sand flies fed No. (%) positive Before treatment.015 Border of lesion (7 97) b 16 5 (31.3) Normal skin (2 74) 6 0 (0) Monocytes (18 58) 4 0 (0) After treatment!.001 Scar (9 63) 1 0 (0) Normal skin (2 40) 0 0 (0) Monocytes (26 69) 3 0 (0) a All patients were exposed to 100 sand flies over the course of 30 min. Only those sand flies that fed and survived were dissected. b P!.05 (Dunn procedure), for the comparison between the no. of flies that fed on the borders of lesions and the no. that fed on normal skin or scars. method used establishes only the presence of parasite DNA, a study of the diagnosis of Chlamydia infection showed that the presence of DNA was associated with the presence of live bacteria and that DNA usually disappeared within 24 h or a maximum of 1 week after bacterial death [41]. Furthermore, in another study, Leishmania DNA levels in blood fell below the limits of detection by PCR in 16 of 17 cases of visceral leishmaniasis within 37 days after the initiation of treatment and, hence, provided an indicator of therapeutic response in visceral leishmaniasis caused by L. chagasi infection [42]. Therefore, the presence of Leishmania kdna in blood and dermal tissue fluid 13 weeks after the initiation of treatment for cutaneous leishmaniasis is likely to indicate the presence of viable parasites. Nevertheless, methods such as real-time PCR targeting RNA should be used to confirm the presence of live parasites [43]. Notably, only 2 of 10 patients in whom Leishmania kdna was detected in skin, scars, or monocytes after treatment had clinical treatment failure. The ED 50 of Sb V for strains isolated before treatment as well as at treatment failure in one of these patients (3565) was 1128 mg Sb V /ml, indicating insensitivity to Sb drugs. Regardless of clinical outcome, parasites that persist after treatment have potential importance in the propagation of drug resistance, given that we have recently documented that standard treatment of American cutaneous leishmaniasis caused by L. (Viannia) infection can indeed select for resistance [12]. Diminished sensitivity to Sb drugs associated with previous treatment has also been documented in dogs; strains isolated from treated dogs were 8 41 times less susceptible to Sb drugs than were those isolated from the same dogs before treatment [44]. The documentation of both primary and secondary resistance in clinical strains [12] is consistent with the persistence and accumulation of parasites with diminished drug sensitivity in endemically exposed human populations. Our results show that L. (Viannia) parasites are present in sites other than the lesion and that they persist in tissues available to sand flies after successful treatment in a substantial proportion of patients. The proportion of positive patients varied by method and sample type: although 11.9% of patients yielded live parasites by either culture of normal skin or xenodiagnosis of lesion border before treatment, 22.8% and 25.9% presented PCR/Southern blot evidence of Leishmania parasites in normal skin or blood monocytes before and after treatment, respectively (table 2). In light of the high number and proportion of infected individuals in areas of endemicity, the plausibility that humans play an important role in the transmission of L. (Viannia) parasites is compelling. The accumulation of observations made over decades indicate that patients with cutaneous leishmaniasis can harbor Leishmania parasites indefinitely and that no currently available treatment completely eliminates them [45]. This fact is particularly relevant to the recent detection of highly anthropophilic Lutzomyia species in urban and periurban areas of Venezuela [46], Brazil [47], Bolivia, and Peru [1, 48, 49]. We believe that the present results, together with these previous findings, call for a reassessment of the role played by humans in transmission cycles of New World cutaneous leishmaniasis, of control strategies, and of the impact that treatment has on transmission. Acknowledgments We thank Maria Ximena Varona, for expert assistance in obtaining dermal tissue fluid aspirates and in parasite isolation; Wilson Cortez, for identifying patients for and enrolling them in this study; and Blanca Es- Leishmania in Skin and Blood JID 2006:194 (15 August) 509

8 corcia, for identification of species and strains by monoclonal-antibody typing. References 1. Campbell-Lendrum D, Dujardin JP, Martinez E, et al. Domestic and peridomestic transmission of American cutaneous leishmaniasis: changing epidemiological patterns present new control opportunities. Mem Inst Oswaldo Cruz 2001; 96: Bejarano EE, Uribe S, Rojas W, Dario Velez I. Phlebotomine sand flies (Diptera: Psychodidae) associated with the appearance of urban leishmaniasis in the city of Sincelejo, Colombia. Mem Inst Oswaldo Cruz 2002; 97: Oliveira CC, Lacerda HG, Martins DR, et al. Changing epidemiology of American cutaneous leishmaniasis (ACL) in Brazil: a disease of the urban-rural interface. Acta Trop 2004; 90: Leonardo FS, Rebêlo JM. A periurbanização de Lutzomyia whitmani em área de foco de leishmaniose cutânea, no Estado do Maranhão, Brasil. Rev Soc Bras Med Trop 2004; 37: Desjeux P. Leishmaniasis: current situation and new perspectives. Comp Immunol Microbiol Infect Dis 2004; 27: Soto J, Toledo J, Vega J, Berman J. Short report: efficacy of pentavalent antimony for treatment of Colombian cutaneous leishmaniasis. Am J Trop Med Hyg 2005; 72: Palacios R, Osorio LE, Grajalew LF, Ochoa MT. Treatment failure in children in a randomized clinical trial with 10 and 20 days of meglumine antimonate for cutaneous leishmaniasis due to Leishmania [Viannia] species. Am J Trop Med Hyg 2001; 64: Romero GA, Guerra MV, Paes MG, Macedo VO. Comparison of cutaneous leishmaniasis due to Leishmania [Viannia] braziliensis and L. [V] guyanensis in Brazil: therapeutic response to meglumine antimoniate. Am J Trop Med Hyg 2001; 65: Alvar J, Canavate C, Gutierrez-Solar B, et al. Leishmania and human immunodeficiency virus coinfection: the first 10 years. Clin Microbiol Rev 1997; 10: Bourreau E, Prevot G, Gardon J, Pradinaud R, Launois P. High intralesional interleukin-10 messenger RNA expression in localized cutaneous leishmaniasis is associated with unresponsiveness to treatment. J Infect Dis 2001; 184: Lira R, Sundar S, Makharia A, et al. Evidence that the high incidence of treatment failures in Indian kala-azar is due to the emergence of antimony-resistant strains of Leishmania donovani. J Infect Dis 1999; 180: Rojas R, Valderrama L, Valderrama M, Varona MX, Ouellette M, Saravia NG. Resistance to antimony and treatment failure in human Leishmania (Viannia) infection. J Infect Dis 2006;193: Travi BL, Montoya J, Solarte Y, Lozano L, Jaramillo C. Studies on the phlebotomine fauna associated with endemic foci in the Pacific Coiast region. Am J Trop Med Hyg 1988; 39: Montoya-Lerma J, Palacios R, Osorio L, Jaramillo C, Cadena H. Further evidence of humans as a source of Leishmania [Viannia] for sand flies. Mem Inst Oswaldo Cruz 1998; 93: Schubach A, Haddad F, Oliveira-Neto MP, et al. Detection of Leishmania DNA by polymerase chain reaction in scars of treated human patients. J Infect Dis 1998; 178: Guevara P, Rojas E, Gonzalez N, et al. Presence of Leishmania braziliensis in blood samples from cured patients or at different stages of immunotherapy. Clin Diagn Lab Immunol 1994; 1: Schubach A, Marzochi MC, Cuzzi-Maya T, et al. Cutaneous scars in American tegumentary leishmaniasis patients: a site of Leishmania [Viannia] braziliensis persistence and viability eleven years after antimonial therapy and clinical cure. Am J Trop Med Hyg 1998; 58: Saravia NG, Weigle K, Segura I, et al. Recurrent lesions in human Leishmania braziliensis infection reactivation or reinfection? Lancet 1990; 336: Ward RD, Lainson R, Shaw JJ. Some methods for membrane feeding of laboratory reared neotropical sandflies (Diptera: Psychodidae). Ann Trop Med Parasitol 1978; 72: Weigle KA, de Davalos M, Heredia P, Molineros R, Saravia NG, D Alessandro A. Diagnosis of cutaneous and mucocutaneous leishmaniasis in Colombia: a comparison of seven methods. Am J Trop Med Hyg 1987; 36: Vergel C, Walker J, Saravia N. Amplification of human DNA by primers targeted to Leishmania kdna and post-genome considerations in diagnostic PCR. Am J Trop Med Hyg 2005; 72: Brujin M, Barker D. Diagnosis of New World leishmaniasis: specific detection of species of the Leishmania braziliensis complex by amplification of kinetoplast DNA. Acta Trop 1992; 52: Weigle K, Labrada LA, Lozano C, Santrich C, Barker D. PCR-based diagnosis of acute and chronic cutaneous leishmaniasis caused by Leishmania [Viannia]. J Clin Microbiol 2002; 40: Saravia NG, Weigle K, Navas C, et al. Heterogeneity, geographic distribution, and pathogenicity of serodemes of Leishmania [Viannia] in Colombia. Am J Trop Med Hyg 2002; 66: Saravia NG, Segura I, Holguin AF, Santrich C, Valderrama L, Ocampo C. Epidemiologic, genetic, and clinical associations among phenotypically distinct populations of Leishmania [Viannia] in Colombia. Am J Trop Med Hyg 1998; 59: Desjeux P. The increase in risk factors for leishmaniasis worldwide. Trans R Soc Trop Med Hyg 2001; 95: Yadon ZE, Rodrigues LC, Davies CR, Quigley MA. Indoor and peridomestic transmission of American cutaneous leishmaniasis in northwestern Argentina: a retrospective case control study. Am J Trop Med Hyg 2003; 68: Costa CH, Gomes RB, Silva MR, et al. Competence of the human host as a reservoir for Leishmania chagasi. J Infect Dis 2000; 182: Rojas E, Scorza JV. Xenodiagnostico con Lutzomyia youngi en casos venozolanos de leishmaniasis cutanea por Leishmania braziliensis. Mem Inst Oswaldo Cruz 1989; 84: Deane L, Rangel E, Paes-Oliveira M, et al. Experimental infection of Lutzomyia longipalpis fed on a patient with cutaneous leishmaniasis due to Leishmania mexicana amazonensis. Mem Inst Oswaldo Cruz 1986; 81: Mendonça MG, de Brito M, Rodrigues E, Bandeira V, Jardim ML, Abath F. Persistence of Leishmania parasites in scars after clinical cure of American cutaneous leishmaniasis: is there a sterile cure? J Infect Dis 2004; 189: Coutinho SG, Pirmez C, Da-Cruz AM. Parasitological and immunological follow-up of American tegumentary leishmaniasis patients. Trans R Soc Trop Med Hyg 2002; 96(Suppl 1):S Weigle KA, Santrich C, Martínez F, et al. Epidemiology of cutaneous leishmaniasis in Colombia: environmental and behavioral risk factors for infection, clinical manifestations, and pathogenicity. J Infect Dis 1993; 168: Courtenay O, Quinnell RJ, Garcez LM, Shaw JJ, Dye C. Infectiousness in a cohort of Brazilian dogs: why culling fails to control visceral leishmaniasis in areas of high transmission. J Infect Dis 2002; 186: Le Fichoux Y, Quaranta JF, Aufeuvre JP, et al. Occurrence of Leishmania infantum parasitemia in asymptomatic blood donors living in an area of endemicity in southern France. J Clin Microbiol 1999; 37: Martín-Sánchez J, Pineda J, Morillas-Márquez F, García-García J, Acedo C, Macías J. Detection of Leishmania infantum kinetoplast DNA in peripheral blood from asymptomatic individuals at risk for parentally transmitted infections: relationship between polymerase chain reaction results and other Leishmania infection markers. Am J Trop Med Hyg 2004; 70: Otero AC, da Silva VO, Luz KG, et al. Short report: occurrence of Leishmania donovani DNA in donated blood from seroreactive Brazilian blood donors. Am J Trop Med Hyg 2000; 62: Travi BL, Tabares CJ, Cadena H, Ferro C, Osorio Y. Canine visceral leishmaniasis in Colombia: relationship between clinical and parasi- 510 JID 2006:194 (15 August) Vergel et al.

9 tologic status and infectivity for sand flies. Am J Trop Med Hyg 2001;64: de Oliveira CI, Báfica A, Oliveira F, Favali CBF, Correa T, Freitas LAR. Clinical utility of polymerase chain reaction based detection of Leishmania in the diagnosis of American cutaneous leishmanisis. Clin Infect Dis 2003; 37:e Marfurt J, Nasereddin A, Niederwieser I, Jaffe CL, Beck HP, Felger I. Identification and differentiation of Leishmania species in clinical samples by PCR amplification of the miniexon sequence and subsequent restriction fragment length polymorphism analysis. J Clin Microbiol 2003; 41: Workowski KA, Lampe MF, Wong KG, Watts MB, Stamm WE. Longterm eradication of Chlamydia trachomatis genital infection after antimicrobial therapy: evidence against persistent infection. JAMA 1993; 270: Disch J, Oliveira MC, Orsini M, Rabello A. Rapid clearance of circulating Leishmania kinetoplast DNA after treatment of visceral leishmaniasis. Acta Tropica 2004; 92: Van der Meide WF, Schoone GJ, Faber WR, et al. Quantitative nucleic acid sequence-based assay as a new molecular tool for detection and quantification of Leishmania parasites in skin biopsy samples. J Clin Microbiol 2005; 43: Lamothe J. Use of meglumine antimoniate in canine leishmaniasis. Vet Rec 2004; 154: Murray HW, Berman JD, Davies CR, Saravia NG. Advances in leishmaniasis. Lancet 2005; 366: Feliciangeli MD. Hourly activity of Lutzomyia ovallesi and L. gomezi (Diptera: Psychodidae), vectors of cutaneous leishmaniasis in northcentral Venezuela. J Med Entomol 1997; 34: Oliveira-Neto MP, Pirmez C, Rangel E, Schubach A, Grimaldi G. An outbreak of American cutaneous leishmaniasis (Leishmania braziliensis braziliensis) in a periurban area of Rio de Janiero city, Brazil: clinical and epidemiological studies. Mem Inst Oswaldo Cruz 1988; 83: Martinez E, Le Pont F, Torrez M, et al. A new focus of cutaneous leishmaniasis due to Leishmania amazonensis in a sub-andean region of Bolivia. Acta Trop 1998; 71: Torrez M, Lopez M, Le Pont F, et al. Lutzomyia nuneztovari anglesi (Diptera: Psychodidae) as a probable vector of Leishmania braziliensis in the Yungas, Bolivia. Acta Trop 1998; 71: Leishmania in Skin and Blood JID 2006:194 (15 August) 511