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Leishmania chagasi infection in cats with dermatologic lesions from an endemic area of visceral leishmaniosis in Brazil
Veterinary Parasitology 178 (2011) 22–28
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Leishmania chagasi infection in cats with dermatologic lesions from an endemic area of visceral leishmaniosis in Brazil Juliana Peloi Vides a , Tatianna Frate Schwardt a , Ludmila Silva Vicente Sobrinho a , Márcia Marinho b , Márcia Dalastra Laurenti c , Alexander Welker Biondo d,1 , Christian Leutenegger e , Mary Marcondes a,∗ a b c d e
Department of Veterinary Clinics, Surgery and Reproduction, São Paulo State University, Arac¸atuba, São Paulo, 16050-680, Brazil Department of Animal Production and Health, São Paulo State University, Arac¸atuba, São Paulo, 16050-680, Brazil Department of Pathology, São Paulo University, São Paulo, 01246-903, Brazil Department of Veterinary Medicine, Federal University of Parana, Curitiba, Parana 80035-050, Brazil IDEXX Laboratories Inc., West Sacramento, CA, 95605, USA
a r t i c l e
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Article history: Received 29 July 2010 Received in revised form 17 December 2010 Accepted 21 December 2010 Keywords: Feline leishmaniosis ELISA IFAT Immunohistochemical RT-PCR
a b s t r a c t Although dogs are considered the main domestic reservoirs for Visceral Leishmaniosis (VL), which is caused in the Americas by Leishmania chagasi, infected cats have also been recently found in endemic areas of several countries and became a public health concern. Accordingly, the purpose of this study was to evaluate cats with dermatologic lesions from an endemic area of VL and the natural infection of L. chagasi. A total of 55 cats were selected between April 2008 and November 2009 from two major animal shelters of Arac¸atuba, Southeastern Brazil. All cats underwent general and dermatologic examinations, followed by direct parasitological examination of lymphoid organs, immunosorbent assay (ELISA) and indirect immunofluorescence (IFAT). In addition, detection of amastigotes was performed by immunohistochemistry (IHC) in skin lesions of all cats. VL was diagnosed in 27/55 (49.1%) cats with dermatological problems. Amastigotes were found in lymphoid organs of 10/27 (37.0%) cats; serology of 14/27 (51.9%), 6/27 (22.2%) and 5/27 (18.5%) cats was positive for ELISA, IFAT and both, respectively. The IHC identified 9/27 (33.3%) cats; 5/27 (18.5%) were positive only for IHC and therefore increased the overall sensitivity. Specific FIV antibodies were found in 6/55 (10.9%) cats, of which 5/6 (83.3%) had leishmaniosis. Real time PCR followed by amplicon sequencing successfully confirmed L. chagasi infection. In conclusion, dermatological lesions in cats from endemic areas was highly associated to visceral leishmaniosis, and therefore skin IHC and differential diagnosis of LV should be always conducted in dermatological patients in such areas. © 2011 Elsevier B.V. All rights reserved.
1. Introduction ∗ Corresponding author at: Departamento de Clínica Cirurgia e Reproduc¸ão Animal, Universidade Estadual Paulista Júlio de Mesquita Filho, Faculdade de Odontologia de Arac¸atuba, Rua Clóvis Pestana, 793. Jardim Dona Amélia, 16050-680, Arac¸atuba, São Paulo, Brazil. Tel.: +55 18 3636 1415, fax: +55 18 3636 1401. E-mail address: [email protected] (M. Marcondes). 1 Department of Veterinary Pathobiology, University of Illinois, IL, 61802, USA. 0304-4017/$ – see front matter © 2011 Elsevier B.V. All rights reserved. doi:10.1016/j.vetpar.2010.12.042
Human Visceral leishmaniosis (VL), endemic in 88 countries and enlisted by the World Health Organization as one of the 17 neglected tropical diseases, is poverty-related and caused by parasitic protozoans of the genus Leishmania (Camargo et al., 2007). The disease has a profound impact on public health with approximately 12 million people yearly infected and only 10% of these new cases
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reported. Dogs are considered the main domestic reservoirs for zoonotic leishmaniosis caused by L. infantum (=L. chagasi), with prevalence ranging from 63% to 80% in endemic areas (Baneth, 2006). However, due to feeding habits of sandflies, other mammalian species may become potential LV reservoirs. The reportedly increasing number of feline leishmaniosis cases suggests that cats may also play a role in the VL epidemiology (Maroli et al., 2007). Since the first reported case of cat leishmaniosis in 1912, from a household where a dog and a child were also infected (Sergent et al., 1912), several cases of feline visceral leishmaniosis have been described in Italy, Brazil, France, Switzerland, Spain, Israel, Portugal and Iran (Ozon et al., 1998; Poli et al., 2002; Rüfenacht et al., 2005; MartínSánchez et al., 2007; Solano-Gallego et al., 2007; Ayllon et al., 2008; Maia et al., 2008; Nasereddin et al., 2008; Costa et al., 2009; Hatam et al., 2009; Sarkari et al., 2009). The first autochthonous case of L. infantum chagasi recorded in the Americas was a domestic cat from São Paulo, Brazil presenting a nose nodular lesion, weight lost and lymphomegaly in 2001 (Savani et al., 2004). Three recent surveillance studies conducted in endemic areas of Southeastern Brazil have shown similar results, with 6/200 (6.5%), 8/200 (4.0%) and 3/52 (5.76%) positive cats in such areas (Rossi, 2007; Costa et al., 2010; Coelho et al., in press), indicating that cats may somehow play a role in the leishmaniosis cycle. The clinical features reportedly associated with Leishmania infection in cats are non-specific and may include ulcer-crusted dermatitis, nodular dermatitis, alopecia, and scaling (Ozon et al., 1998; Pennisi et al., 2004; Nasereddin et al., 2008). Visceral forms of disease have also been described in cats and include weight loss, dehydration, lymphadenopathy, lymphomegaly and hepatic, splenic, respiratory and digestive disorders (Ozon et al., 1998; Hervás et al., 1999; Pennisi et al., 2004; Grevot et al., 2005; Leiva et al., 2005; Vita et al., 2005; Ayllon et al., 2008; Nasereddin et al., 2008; Silva et al., 2010). Although not pathognomonic, dermatological lesions may be important for screening infected animals in endemic areas. Accordingly, the purpose of this study was to evaluate cats with dermatologic lesions from an endemic area of VL and its association to the natural infection of L. chagasi. 2. Material and methods 2.1. Study subjects All cats presenting dermatological lesions from April, 2008 to November, 2009 at two major shelters located in Arac¸atuba, São Paulo, Brazil were selected for the present study. All cats underwent general and dermatologic examinations, followed by diagnosis of VL based on the direct observation of L. sp. amastigote forms in bone marrow, lymph nodes, spleen and liver smears as well as by ELISA, IFAT and additionally immunohistochemistry of skin lesions. The present study has been approved by the Ethics Committee in Animal Experimentation and Animal Welfare of São Paulo State University (UNESP) Arac¸atuba (protocol number 2008-005698).
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2.2. Parasitological diagnosis Direct parasitological diagnosis was performed by light microscopy in bone marrow, lymph nodes, spleen and liver smears obtained by fine needle aspiration cytology on hematological slides stained by Diff-Quick (Panótico Rápido® , Laborclin, São Paulo, Brazil).
2.3. Enzyme-Linked Immunosorbent Assay (ELISA) for Leishmania sp In each well of ELISA plates, 100 L of L. chagasi (MHOM/BR/72) total promastigote lysate (20 g/mL), previously diluted in sodium carbonate–bicarbonate buffer (0.05 M, pH 9.6), was added. After overnight incubation at 4 ◦ C, plates were washed with phosphate buffered saline (PBS) containing 0.05% Tween-20 and were blocked for two hours at 37 ◦ C using 150 L of 10% skimmed milk (Molico® , Nestlé, São Paulo, Brazil). Then 100 L of sera diluted (1:200) in PBS Tween-20 was added to each well and incubated for 60 min at 37 ◦ C. After a new wash, 100 L of protein A labeled with peroxidase (Sigma–Aldrich–P8651) was added to each well and incubated for 45 min at 37 ◦ C. The enzyme reaction was carried out with 100 L of tetramethylbenzidine (TMB) (BD–555214) solution. Reaction was interrupted adding 50 L of H2 SO4 1 M and the optical density (OD) read at 450 nm using an ELISA reader (Labsystems Multiskan EX, Flow Laboratories International SA, Lugano, Switzerland). Negative and positive controls were included in each plate in duplicate, and values were expressed by serum OD mean. Cut-off values were determined by analysis of serum samples from 50 healthy cats from a non-endemic area for leishmaniosis. The mean and standard deviation (S.D.) were calculated. The mean value plus 3 S.D. was considered as the cut-off point.
2.4. Indirect Immunofluorescente Antibody Test (IFAT) for L. sp. Slides with twelve previously marked circles were sensitized by adding to each circle 20 L suspension of parasites (MHOM/BR/72) in buffered saline solution (PBS) at a concentration of 8.1 × 106 promastigotes/mL. Twenty micrograms of serum samples of cats to be tested were then added to each circle. A positive and a negative control sample, diluted at concentrations of 1:20 and 1:40 in PBS, were added in each slide. The slides were then incubated at 37 ◦ C for 30 min and washed by immersion in three baths, for five minutes each, in PBS. After drying, 20 L of an anti-cat IgG FITC conjugate (F4262, Sigma–Aldrich, Saint Louis, MO, USA), diluted at 1:100 in PBS containing 4 mg% of Evan’s Blue was added in each circle. Slides were washed again in PBS, covered with buffered glycerin and a cover slip, and then examined on a fluorescent microscope. For positive serum samples, parasites displayed a bright-green peripheral stain with a dull fluorescence of the cytoplasm. Positive sera were serially diluted and tested to establish the maximum reaction titer.
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2.5. Immunohistochemistry (IHC) Skin biopsies taken from lesioned area by 5-mm punch were fixed in 10% neutral-buffered formalin solution for 72 h. The tissues samples were dehydrated, cleared, embedded in paraffin, cut (4–5 mm), and stained with hematoxylin and eosin (HE). Deparaffinized slides were hydrated and incubated in 2% hydrogen peroxide 10 (v/v) in 0.01 M PBS, pH 6.0, to block endogenous peroxidase activity, followed by incubation with 10% skimmed milk to block nonspecific immunoglobulin absorption to tissues. A heterologous hyperimmune serum from mouse infected with L. amazonensis was used as primary antibody. Slides were incubated for 18 h at 4 ◦ C in a humid chamber. After washing in PBS, the slides were incubated with biotinylated side antibody (DAKO), washed in PBS again, and the incubated with the streptavidin–peroxidase complex (DAKO) for 45 min each at 37 ◦ C. The reaction was developed with a diaminobenzidine (K3468, DAKO,) solution and hydrogen peroxide 10 (v/v). Finally, slides were dehydrated, cleared, counter-satined with Harris Hematoxylin, and mounted with coverlips. 2.6. Detection of FeLV antigen and FIV antibody To evaluate the impact of immunosuppressive retroviruses, all serum samples were tested for FeLV antigen (p27) and FIV antibody using a commercial rapid assay kit (SNAP® FIV Antibody/FeLV Antigen Combo Test: IDEXX Laboratories, Westbrook, ME, USA). 2.7. Real time polymerase chain reaction and sequencing To ensure that infection was caused by L. chagasi, EDTA anticoagulated whole blood samples were randomly obtained from positive cats, stored at −20 ◦ C and DNA extracted under standard protocols on a commercial platform (Corbett XTractor-Gene, Qiagen, Valencia, CA, USA). A housekeeping gene (18S rRNA) was used to determine DNA content and quality. The PCR test was based on IDEXX’s proprietary real-time PCR oligonucleotides (IDEXX Laboratories, Westbrook, ME, USA). Briefly, highly conserved metalloproteinase gp63 gene sequences were aligned and a region selected for 2 primers and a hydrolysis probe designed using commercial software (PrimerExpress Version 3.0, Applied Biosystems, Foster City, CA, USA). An additional 2 primers flanking the real-time PCR assay were designed for sequence verification purposes. Real-time PCR was run with standard primer and probe concentrations (ABI) using a commercially available mastermix (Roche LightCycler® 480 Probes Master, Roche Applied Science, Indianapolis, IN, USA). Real-time PCR was performed using default cycling conditions on a commercial instrument (Roche LC480 in the 384-well plate configuration, Roche Applied Science, Indianapolis IN, USA). Amplicons were submitted for sequencing according to standard protocols. 2.8. Statistical analysis Kappa test was used to verify the diagnostic test agreement. Differences were considered significant when the P value was <0.05. Statistical analysis was performed by a
Table 1 Results of the direct parasitological examination of the lymphoid organs (popliteal lymph nodes–PL, bone marrow–BM, spleen–S and liver–L); immunohistochemical reaction (IHQ) of skin lesions; ELISA and indirect immunofluerecence reaction (IFAT) for visceral leishmaniasis; and FIV specific antibodies detection in 27 cats naturally infected by L. chagasi living in an endemic area for visceral leishmaniasis in Brazil. Animal
Parasitological examination
ELISA
IFAT
IHQ
FIV
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27
+BM – +BM/ +S/ +L – – +PL/ +L – – – +BM/ +S +BM/ +S +PL/ +S +BM – – – +BM – +BM +PL – – – – – – –
− − − + + + + − + − + − − + + − − − − − + + + − + + +
− − − − + + + − − − − − + − − − − − − − − − − − + + −
− + + − − + − + − − − − − − + + − + − − − − − + + − −
− − − − − + − − − − − − − − − − − − − − − + + − + − −
commercially available software program (InStat® , GraphPad Software Inc., La Jolla, CA, USA).
3. Results 3.1. Study subjects A total of 55 cats presenting dermatological lesions, 17 males and 10 females with age average of 3.5 years old, were obtained from April, 2008 to November, 2009. From the total number of evaluated animals, 27/55 (49.1%) were positive for visceral leishmaniosis by at least one of the diagnostic methods applied.
3.2. Parasitological diagnosis Amastigote forms of L. sp. were identified in lymphoid organs from 10/55 (18.2%) infected cats (Table 1). The parasite was more frequently found in bone marrow (70.0%), followed by spleen (40.0%), lymph nodes (30.0%) and liver (20.0%). In five cats (50.0%) the parasite was identified in more than one organ. Three positive cats on parasitological test were also positive for ELISA, two for IFAT and one animal was considered positive by both serological techniques.
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3.3. ELISA and IFAT for L. sp. The calculated cut-off value used in the present study was 0.2765 for ELISA and 1:40 for IFAT. The seroprevalence of leishmaniosis found was 14/55 (25.4%) cats by ELISA and 6/55 (10.9%) cats by IFAT (Table 1). Levels of L. sp. antibodies in ELISA positive cats ranged from 0.278 to 1.631, with a mean of 0.710. Five IFAT positive cats had titers of 1:40 and one 1:80. From these six animals, five were also positive by ELISA. Amastigotes of L. sp were identified in lymphoid organs of 4/14 ELISA and 3/6 IFAT positive cats. A poor kappa overall agreement was found between serological tests and parasitological diagnosis ( = −0.006 and = 0.135, respectively) 3.4. Immunohistochemistry (IHC) The anti-Leishmania immunohistochemical assay allowed the identification of nine (16.4%) positive animals (Table 1). The parasite was identified on the tip of the pinna (55.5%), bridge of the nose (22.2%), chin area (11.1%) and around the eyes (11.1%). In five cats, although immunostaining of lesioned skin areas was positive, parasites were not identified in lymphoid organs and their serological tests resulted negative. 3.5. Detection of FeLV antigen and FIV antibody FIV specific antibodies were found in 6/55 cats (10.9%), from which 5/6 (83.3%) were Leishmania infected (Table 1). Amastigotes were evidenced in 3/5 (60.0%) cats with coinfection and all of them had anti-L. chagasi IgG antibodies, showing an association between FIV and L. chagasi specific antibodies. FeLV antigens were not detected in any cat.
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evident dermatological sign was the presence of alopecia observed in the cephalic region, particularly the pinna of 26 animals. Alopecia was also observed in the limbs, dorsal trunk, abdomen and tail. Ulcers with hemorrhagic crusts were observed in nine animals, eight of them simultaneously on the pinna and bridge of the nose. One animal initially presented alopecia and crusts only in the left pinna; after 60 days the cat tested positive and dermatological condition had evolved to deep ulcers with hemorrhagic crusts (Fig. 1). Erythema was found in eight cats on the pinna, abdomen, limbs, cervical area and bridge of the nose. One of these animals had a nummular erythematous alopecia measuring one centimeter in diameter on the bridge of the nose (Fig. 2), with positive fungal culture for Microsporum canis. The immunohistochemistry reaction of this area showed immunostaining for amastigote of L. sp. Hyperpigmentation, hyperkeratosis, lichenification and exudation on dorsal trunk in cephalic and abdominal region was observed in three animals (Fig. 2). One cat presented scaling, comedones and epidermal collarettes on dorsal trunk and pinna, characterizing a seborrheic dermatitis condition. A total of 19/27 (70.4%) Leishmania infected cats had systemic disorders along dermatological lesions, including lymphadenopathy (52.6%), dehydration (52.6%), weight loss (36.8%), diarrhea (21%), bilateral mucopurulent ocular discharge (15.8%), changes in the state of consciousness (15.8%), ulcerations in the oral cavity (10.5%), dyspnea (5.3%), bilateral purulent nasal discharge (5.3%) and corneal opacity (5.3%). Surprisingly, 8/27 (29.6%) cats had no clinical manifestation of systemic disease besides the dermatological lesions.
3.6. Real time polymerase chain reaction
4. Discussion
Real-time PCR was applied in samples from three cats. Average analytical sensitivity was ten molecules per reaction (reproducible in triplicate analysis). Analytical specificity was confirmed by sequencing a positive real-time PCR reaction. Analytical specificity was further confirmed by testing DNA extracted from 23 bacterial, protozoal and rickettsial isolates. Reproducibility (CV on CT values and on absolute quantified numbers), amplification efficiency, linearity, square values and signal to noise ratio parameters all passed the required criteria and thresholds. Nucleotide sequence obtained resulted in an identity level between 97% and 100% with XM 001463664.1, XM 001463663.1, XM 001463662.1, XM 001463661.1, AM 502228.1 sequences (geneBank), confirming L. chagasi diagnosis.
The outcome of 27/55 (49.1%) positive cats for Leishmania was significantly higher than previous studies in the same Brazilian endemic area, which have found 6/200 (6.5%), 8/200 (4.0%) and 3/52 (5.76%) positive cats (Rossi, 2007; Costa et al., 2010; Coelho et al., in press). The screening of cats with skin problems in our study caused a 7.5 to 12.3-fold increase in positivity, clearly showing the importance of dermatological signs for diagnosis of Leishmania. Our data was higher even when compared to highly endemic areas of other countries, such as 7/23 (30.4%) cats found in Portugal (Maia et al., 2008). In our study, amastigote forms of L. sp. was found in lymphoid organs of 10/55 (18.2%) cats, which is around 4.5fold higher than previously found in two cat surveys, with 8/200 (4.0%) (Costa et al., 2010) and 2/52 (3.84%) (Coelho et al., in press) positive samples. A reasonable explanation for this finding would be that once amastigotes have produced skin lesions, lymphoid tissues have probably been invaded as well. Nevertheless, skin problem screening has greatly increased Leishmania detection in cats. Divergent results were observed in amastigotes found in bone marrow (7/10; 70.0%) and lymph nodes (3/10; 30.0%) of cats with dermatological lesions when compared to bone marrow (3/8; 37.5%) and lymph nodes (7/8; 87.5%) in a general cat population previously studied (Costa et al., 2010).
3.7. Clinical signs Based on the topographic distribution of lesions, clinical signs included 15/27 (55.5%) cats with cephalic involvement, 6/27 (22.2%) with lesions on limbs, 6/27 (22.2%) on dorsal trunk, 3/27 (11.1%) on the tail and 3/27 (11.1%) on the abdomen. The most affected areas in the cephalic region were the pinna (60.0%), followed by generalized involvement (20.0%) and the bridge of the nose (20.0%). The most
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Fig. 1. Skin lesions in cats naturally infected by L. chagasi. (a) Alopecia on the external surface of the left pinna with erythema and crusts (b); the same animal showing, after 60 days, deep ulceration on the tip of the left pinna with hemorrhagic crusts (c); alopecia with active bleeding and hemorrhagic crusts on the base of the right pinna (d); the same animal, demonstrating that the lesions were bilateral and symmetric.
No reasonable cause has been found to explain the more likelihood of bone marrow for parasites to be found in the present study and therefore a higher sensitivity when compared to other lymphoid tissues. Interestingly, five cats had the diagnosis confirmed by just one of the four evaluated organs, which emphasizes the importance of simultaneous parasite searching for optimum sensitivity. Seroprevalence observed in the present study was significantly higher than previously found in randomly selected cats from the same area, which showed prevalence of 3% and 11.5% (Rossi, 2007; Costa et al., 2009). ELISA has been positive in 14/55 (25.5%) cats and IFAT in 6/55 (10.9%) cats, of which 5/6 were positive for both IFAT and ELISA. Levels of antibodies in ELISA positive cats ranged between 0.278 and 1.631, reaching up to 6-fold the cut-off in some cats as previously observed (Solano-Gallego et al., 2007).
Since serum samples of known negative cats were still reactive at a dilution of 1:20, the cut-off used to identify positive animals was 1:40, identical to previously obtained by others (Silva et al., 2008; Duarte et al., 2010). In the present study, IFAT positive cats all had low titers, varying between 1:40 (5/6) and 1:80 (1/6), differing from previous results of titers up to 1:320 in L. infantum naturally infected cats (Vita et al., 2005; Silva et al., 2008). The low agreements between the parasitological method and ELISA ( = −0.006), and the parasitological method and IFAT ( = 0.1356) were similar to those previously obtained (Rossi, 2007). Inconsistency when comparing results may be clearly explained since seven infected cats were only identified by ELISA, whereas only 4/14 ELISA positive cats had shown parasites in lymphoid organs. On the other hand, 3/6 IFAT positive cats
Fig. 2. Skin lesions in cats naturally infected by L. chagasi. (a) Alopecic, erythematous and circular area, of approximately one centimeter of diameter, in which amastigotes of the parasite were identified (b); cervical area of another cat exhibiting alopecia, erythema and exudation.
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had amastigotes detected on direct smears from lymphoid organs, in accordance of a recent study in which 91.3% of serologically reactive animals were parasitologically negative (Costa et al., 2010). Due to its low sensitivity, traditional histological diagnosis of leishmaniosis may be unsuccessful to detect the parasite. Immunohistochemical technique (IHC), on the other hand, is a highly sensitive and specific tool for the diagnosis of visceral leishmaniosis (Ferrer et al., 1988; Bourdoiseau et al., 1997). In the present study, the IHC for detection of L. sp. on the skin has increased the rate of positive results compared to the other performed methods. Amastigotes were easily observed by IHC in nine cats, five of them positive only by IHC, confirming previous descriptions in which the use of this exam may increase the possibility of finding parasites even when in low amounts in the evaluated tissues (Ferrer et al., 1988; Bourdoiseau et al., 1997). Skin parasitism may post an additional public health concern due to higher potential of infection to sand flies, since sand flies may be infected by L. infantum when feeding on a cat with visceral leishmaniosis, as recently demonstrated by xenodiagnosis (Maroli et al., 2007). In the present study, asymptomatic presentation was more frequent than symptomatic, which may lead public health professionals to underestimate the actual number of infected cats. This particular feature may be consequence of differences in immunological response of cats when compared to dogs. While dogs mainly present a humoral response, cats appear to have a higher degree of natural resistance due to a more cellular immune response (Solano-Gallego et al., 2007). In cases of immunosuppressive events, such as retrovirus infection, compromised cellular immunity may lead to the dissemination and visceralization of the parasite (Martín-Sánchez et al., 2007). Accordingly, 6/55 (10.9%) cats had anti-Feline Immunodeficiency Virus (FIV) antibodies, from which five (83.3%) had leishmaniosis, demonstrating an association between immunosuppression on FIV and Leishmania coinfection. These findings are in agreement with a previous study where a high L. infantum seropositivity rate in FIV-positive cats was reported (Pennisi et al., 2004). Since no FeLV antigens were detected in the present study, we were not able to determine whether FeLV infection is a potential risk factor for feline L. infantum infection as previously observed (Sherry et al., 2010). Alopecia, particularly observed on the pinnae, deep ulcerations and scaling were the most common dermatologic signs observed in this study. Some infected cats had only erythematous areas with alopecia or crusts without any nodules or ulcerated areas. These skin disorders may be associated with other dermatological conditions frequently affecting cats (Navarro et al., 2010). Consequently, leishmaniosis may be underestimated during clinical evaluation and differential diagnosis and clinicians should be aware of the concomitant presentation of dermatological problems, particularly when working in endemic areas. As previously observed, a total of 19/27 (70.4%) infected cats also showed systemic disorders such as lymphadenopathy, weight loss, mucopurulent ocular discharge, bilateral corneal opacity, purulent nasal discharge,
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changes in consciousness state, oral cavity ulcers, dyspnea and diarrhea (Poli et al., 2002; Vita et al., 2005). Since the aim of the study was strictly apply Real-time PCR to insure Leishmania species rather than a diagnostic approach, tests were performed only in three positive cats. L. chagasi strains were consistently identified by sequence analysis in all of them. Although two cases of L. amazonensis infection have been previously reported in domestic dogs living in the same area (Tolezano et al., 2007), two PCR positive cats were from the same shelter where 17 infected animals were found, and the third one was from other shelter were 10 infected animals were detected. However, further studies should be conducted in order to fully establish the Leishmania distribution in cat populations of endemic areas, as well as the cat role in the disease cycle. 5. Conclusion Based on the high association to dermatological lesions, leishmaniosis should be included as differential diagnosis of skin diseases of cats living in endemic areas. Moreover, immunohistochemistry should be always employed to increase diagnostic sensitivity. Finally, cats may represent a source of infection to sand flies as they have high skin parasitism. Acknowledgment We are grateful to the Fundac¸ão de Amparo à Pesquisa do Estado de São Paulo (FAPESP), project 2009/52812-3 for financial support. References Ayllon, T., Tesouro, M.A., Amusategui, I., Villaescusa, A., Rodriguez-Franco, F., Sainz, A., 2008. Serologic and Molecular evaluation of Leishmania infantum in cats from Central Spain. Ann. N. Y. Acad. Sci. 1149, 361–364. Baneth, G., 2006. Leishmaniasis. In Greene, C.E. (Ed.), Infectious diseases. Elsevier, Canadá, pp. 685–698. Bourdoiseau, G., Marchal, T., Magnol, J.P., 1997. Immunohistochemical detection of Leishmania infantum in formalin-fixed, paraffinembedded sections of canine skin and lymph nodes. J. Vet. Diag. Invest. 9, 439–440. Camargo, J.B., Troncarelli, M.Z., Ribeiro, M.G., Langoni, H., 2007. Canine visceral leishmaniosis: aspects of public health and control. Clín. Vet. 12, 86–92. Coelho, W.M.D., Pereira, V.B.R., Langoni, H., Bresciani, K.D.S. Molecular detection of Leishmania sp. in cats (Felis catus) from Andradina Municipality, São Paulo State, Brazil. Vet. Parasitol., in press. Costa, T.A.C., Rossi, C.N., Laurenti, M.D., Gomes, A.A.D., Vides, J.P., Sobrinho, L.S.V., Marcondes, M., 2010. Occurrence of leishmaniosis in cats of an endemic area for visceral leishmaniosis. Braz. J. Vet. Anim. Sci. 47, 213–217. Costa, T.A.C., Rossi, C.N., Laurenti, M.D., Gomes, A.A.D., Vides, J.P., Sobrinho, L.S.V., Costa, D.C., Marcondes, M., 2009. Use of enzyme-linked immunosorbent assay for serological diagnosis of Leishmaniasis in cats. In: Proceedings of the 34th WSAVA Congress , São Paulo, Brazil, pp. 93–94. Duarte, A., Castro, I., da Fonseca, I.M.P., Almeida, V., de Carvalho, L.M.M., Meireles, J., Fazendeiro, M.I., Tavares, L., Vaz, Y., 2010. Survey of infectious and parasitic diseases in stray cats at the Lisbon Metropolitan Area. Portugal. J. Feline Med. Surg. 12, 441–446. Ferrer, L., Rabanal, R., Fondevila, D., Ramos, J.A., Domingo, M., 1988. Skin lesions in canine leishmaniasis. J. Small Ann. Pract. 29, 381–388. Grevot, A., Jaussaud Hugues, P., Marty, P., Pratlong, F., Ozon, C., Haas, P., Breton, C., Bourdoiseau, G., 2005. Leishmaniosis due to Leishmania infantum in a FIV and FeLV positive cat with a squamous cell
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Rüfenacht, S., Sager, H., Müller, N., Schaerer, V., Heier, A., Welle, M.M., Roosje, P.J., 2005. Two cases of feline leishmaniosis in Switzerland. Vet. Rec. 23, 542–545. Sarkari, B., Hatam, G.R., Adnani, S.J., Asgari, Q., 2009. Seroprevalence of feline leishmaniasis in areas of Iran where Leishmania infantum is endemic. Ann. Trop. Med. Parasitol. 103, 275–277. Savani, E.S.M.M., de Oliveira Camargo, M.C.G., de Carvalho, M.R., Zampieri, R.A., dos Santos, M.G., D’áuria, S.R.N., Shaw, J.J., Floeter-Winter, L.M., 2004. The first record in the Americas of an autochthonous case of Leishmania (Leishmania) infantum chagasi in a domestic cat (Felix catus) from Cotia County, São Paulo State Brazil. Vet. Parasitol. 25, 229–233. Sergent, Ed., Sergent, Et., Lombard, J, Quilichini, M., 1912. Leishmaniosis in Algeria, with simultaneous infection of a child, a dog and a cat in the same household. Bull. Soc. Pathol. Exot. 5, 93–98. Sherry, K., Miró, G., Trotta, M., Miranda, C., Montoya, A., Espinosa, C., Ribas, F., Furlanello, T., Solano-Gallego, S., 2010. A serological and molecular study of Leishmania infantum infection in cats from the Island of Ibiza (Spain). Vector Borne and Zoonotic Dis. (Aug 30) (epub ahead of print). Silva, A.V.M., Cândido, C.D.S., Pereira, D.P., Brazil, R.P., Carreira, C.A., 2008. The first record of American visceral leishmaniasis in domestic cats from Rio de Janeiro, Brazil. Acta Trop. 105, 92–94. Silva, S.M., Rabelo, P.F.-B., Gontijo, N.F., Ribeiro, R.R., Melo, M.N., Ribeiro, V.M., Michalick, M.S.M., 2010. First reporto f infection of Lutzomyia longipalpis by Leishmania (Leishmania) infantum from a naturally infected cat of Brazil. Vet. Parasitol. 174, 150–154. Solano-Gallego, L., Rodríguez-Cortés, A., Iniesta, L., Quintana, J., Pastor, J., Espada, Y., Portús, M., Alberola, J., 2007. Cross-sectional serosurvey of feline leishmaniasis in ecoregions around the Northwestern Mediterranean. Am. J. Trop. Med. Hyg. 76, 676–680. Tolezano, J.E., Uliana, S.R.B., Taniguchi, H.H., Araujo, M.F.L., Barbosa, J.A.R., Barbosa, J.E.R., Floeter-Winter, L.M., Shaw, J.J., 2007. The first records of Leishmania (Leishmania) amazonensis in dogs (Canis familiaris) diagnosed clinically as having canine visceral leishmaniasis from Arac¸atuba County, São Paulo State. Brazil. Vet. Parasitol. 149, 280–284. Vita, S., Santori, D., Aguzzi, I., Petrotta, E., Luciani, A., 2005. Feline leishmaniasis and ehrlichiosis: serological investigation in Abruzzo region. Vet. Res. Commun. 29, 319–321.
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Leishmania chagasi infection in cats with dermatologic lesions from an endemic area of visceral leishmaniosis in Brazil Juliana Peloi Vides a , Tatianna Frate Schwardt a , Ludmila Silva Vicente Sobrinho a , Márcia Marinho b , Márcia Dalastra Laurenti c , Alexander Welker Biondo d,1 , Christian Leutenegger e , Mary Marcondes a,∗ a b c d e
Department of Veterinary Clinics, Surgery and Reproduction, São Paulo State University, Arac¸atuba, São Paulo, 16050-680, Brazil Department of Animal Production and Health, São Paulo State University, Arac¸atuba, São Paulo, 16050-680, Brazil Department of Pathology, São Paulo University, São Paulo, 01246-903, Brazil Department of Veterinary Medicine, Federal University of Parana, Curitiba, Parana 80035-050, Brazil IDEXX Laboratories Inc., West Sacramento, CA, 95605, USA
a r t i c l e
i n f o
Article history: Received 29 July 2010 Received in revised form 17 December 2010 Accepted 21 December 2010 Keywords: Feline leishmaniosis ELISA IFAT Immunohistochemical RT-PCR
a b s t r a c t Although dogs are considered the main domestic reservoirs for Visceral Leishmaniosis (VL), which is caused in the Americas by Leishmania chagasi, infected cats have also been recently found in endemic areas of several countries and became a public health concern. Accordingly, the purpose of this study was to evaluate cats with dermatologic lesions from an endemic area of VL and the natural infection of L. chagasi. A total of 55 cats were selected between April 2008 and November 2009 from two major animal shelters of Arac¸atuba, Southeastern Brazil. All cats underwent general and dermatologic examinations, followed by direct parasitological examination of lymphoid organs, immunosorbent assay (ELISA) and indirect immunofluorescence (IFAT). In addition, detection of amastigotes was performed by immunohistochemistry (IHC) in skin lesions of all cats. VL was diagnosed in 27/55 (49.1%) cats with dermatological problems. Amastigotes were found in lymphoid organs of 10/27 (37.0%) cats; serology of 14/27 (51.9%), 6/27 (22.2%) and 5/27 (18.5%) cats was positive for ELISA, IFAT and both, respectively. The IHC identified 9/27 (33.3%) cats; 5/27 (18.5%) were positive only for IHC and therefore increased the overall sensitivity. Specific FIV antibodies were found in 6/55 (10.9%) cats, of which 5/6 (83.3%) had leishmaniosis. Real time PCR followed by amplicon sequencing successfully confirmed L. chagasi infection. In conclusion, dermatological lesions in cats from endemic areas was highly associated to visceral leishmaniosis, and therefore skin IHC and differential diagnosis of LV should be always conducted in dermatological patients in such areas. © 2011 Elsevier B.V. All rights reserved.
1. Introduction ∗ Corresponding author at: Departamento de Clínica Cirurgia e Reproduc¸ão Animal, Universidade Estadual Paulista Júlio de Mesquita Filho, Faculdade de Odontologia de Arac¸atuba, Rua Clóvis Pestana, 793. Jardim Dona Amélia, 16050-680, Arac¸atuba, São Paulo, Brazil. Tel.: +55 18 3636 1415, fax: +55 18 3636 1401. E-mail address: [email protected] (M. Marcondes). 1 Department of Veterinary Pathobiology, University of Illinois, IL, 61802, USA. 0304-4017/$ – see front matter © 2011 Elsevier B.V. All rights reserved. doi:10.1016/j.vetpar.2010.12.042
Human Visceral leishmaniosis (VL), endemic in 88 countries and enlisted by the World Health Organization as one of the 17 neglected tropical diseases, is poverty-related and caused by parasitic protozoans of the genus Leishmania (Camargo et al., 2007). The disease has a profound impact on public health with approximately 12 million people yearly infected and only 10% of these new cases
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reported. Dogs are considered the main domestic reservoirs for zoonotic leishmaniosis caused by L. infantum (=L. chagasi), with prevalence ranging from 63% to 80% in endemic areas (Baneth, 2006). However, due to feeding habits of sandflies, other mammalian species may become potential LV reservoirs. The reportedly increasing number of feline leishmaniosis cases suggests that cats may also play a role in the VL epidemiology (Maroli et al., 2007). Since the first reported case of cat leishmaniosis in 1912, from a household where a dog and a child were also infected (Sergent et al., 1912), several cases of feline visceral leishmaniosis have been described in Italy, Brazil, France, Switzerland, Spain, Israel, Portugal and Iran (Ozon et al., 1998; Poli et al., 2002; Rüfenacht et al., 2005; MartínSánchez et al., 2007; Solano-Gallego et al., 2007; Ayllon et al., 2008; Maia et al., 2008; Nasereddin et al., 2008; Costa et al., 2009; Hatam et al., 2009; Sarkari et al., 2009). The first autochthonous case of L. infantum chagasi recorded in the Americas was a domestic cat from São Paulo, Brazil presenting a nose nodular lesion, weight lost and lymphomegaly in 2001 (Savani et al., 2004). Three recent surveillance studies conducted in endemic areas of Southeastern Brazil have shown similar results, with 6/200 (6.5%), 8/200 (4.0%) and 3/52 (5.76%) positive cats in such areas (Rossi, 2007; Costa et al., 2010; Coelho et al., in press), indicating that cats may somehow play a role in the leishmaniosis cycle. The clinical features reportedly associated with Leishmania infection in cats are non-specific and may include ulcer-crusted dermatitis, nodular dermatitis, alopecia, and scaling (Ozon et al., 1998; Pennisi et al., 2004; Nasereddin et al., 2008). Visceral forms of disease have also been described in cats and include weight loss, dehydration, lymphadenopathy, lymphomegaly and hepatic, splenic, respiratory and digestive disorders (Ozon et al., 1998; Hervás et al., 1999; Pennisi et al., 2004; Grevot et al., 2005; Leiva et al., 2005; Vita et al., 2005; Ayllon et al., 2008; Nasereddin et al., 2008; Silva et al., 2010). Although not pathognomonic, dermatological lesions may be important for screening infected animals in endemic areas. Accordingly, the purpose of this study was to evaluate cats with dermatologic lesions from an endemic area of VL and its association to the natural infection of L. chagasi. 2. Material and methods 2.1. Study subjects All cats presenting dermatological lesions from April, 2008 to November, 2009 at two major shelters located in Arac¸atuba, São Paulo, Brazil were selected for the present study. All cats underwent general and dermatologic examinations, followed by diagnosis of VL based on the direct observation of L. sp. amastigote forms in bone marrow, lymph nodes, spleen and liver smears as well as by ELISA, IFAT and additionally immunohistochemistry of skin lesions. The present study has been approved by the Ethics Committee in Animal Experimentation and Animal Welfare of São Paulo State University (UNESP) Arac¸atuba (protocol number 2008-005698).
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2.2. Parasitological diagnosis Direct parasitological diagnosis was performed by light microscopy in bone marrow, lymph nodes, spleen and liver smears obtained by fine needle aspiration cytology on hematological slides stained by Diff-Quick (Panótico Rápido® , Laborclin, São Paulo, Brazil).
2.3. Enzyme-Linked Immunosorbent Assay (ELISA) for Leishmania sp In each well of ELISA plates, 100 L of L. chagasi (MHOM/BR/72) total promastigote lysate (20 g/mL), previously diluted in sodium carbonate–bicarbonate buffer (0.05 M, pH 9.6), was added. After overnight incubation at 4 ◦ C, plates were washed with phosphate buffered saline (PBS) containing 0.05% Tween-20 and were blocked for two hours at 37 ◦ C using 150 L of 10% skimmed milk (Molico® , Nestlé, São Paulo, Brazil). Then 100 L of sera diluted (1:200) in PBS Tween-20 was added to each well and incubated for 60 min at 37 ◦ C. After a new wash, 100 L of protein A labeled with peroxidase (Sigma–Aldrich–P8651) was added to each well and incubated for 45 min at 37 ◦ C. The enzyme reaction was carried out with 100 L of tetramethylbenzidine (TMB) (BD–555214) solution. Reaction was interrupted adding 50 L of H2 SO4 1 M and the optical density (OD) read at 450 nm using an ELISA reader (Labsystems Multiskan EX, Flow Laboratories International SA, Lugano, Switzerland). Negative and positive controls were included in each plate in duplicate, and values were expressed by serum OD mean. Cut-off values were determined by analysis of serum samples from 50 healthy cats from a non-endemic area for leishmaniosis. The mean and standard deviation (S.D.) were calculated. The mean value plus 3 S.D. was considered as the cut-off point.
2.4. Indirect Immunofluorescente Antibody Test (IFAT) for L. sp. Slides with twelve previously marked circles were sensitized by adding to each circle 20 L suspension of parasites (MHOM/BR/72) in buffered saline solution (PBS) at a concentration of 8.1 × 106 promastigotes/mL. Twenty micrograms of serum samples of cats to be tested were then added to each circle. A positive and a negative control sample, diluted at concentrations of 1:20 and 1:40 in PBS, were added in each slide. The slides were then incubated at 37 ◦ C for 30 min and washed by immersion in three baths, for five minutes each, in PBS. After drying, 20 L of an anti-cat IgG FITC conjugate (F4262, Sigma–Aldrich, Saint Louis, MO, USA), diluted at 1:100 in PBS containing 4 mg% of Evan’s Blue was added in each circle. Slides were washed again in PBS, covered with buffered glycerin and a cover slip, and then examined on a fluorescent microscope. For positive serum samples, parasites displayed a bright-green peripheral stain with a dull fluorescence of the cytoplasm. Positive sera were serially diluted and tested to establish the maximum reaction titer.
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2.5. Immunohistochemistry (IHC) Skin biopsies taken from lesioned area by 5-mm punch were fixed in 10% neutral-buffered formalin solution for 72 h. The tissues samples were dehydrated, cleared, embedded in paraffin, cut (4–5 mm), and stained with hematoxylin and eosin (HE). Deparaffinized slides were hydrated and incubated in 2% hydrogen peroxide 10 (v/v) in 0.01 M PBS, pH 6.0, to block endogenous peroxidase activity, followed by incubation with 10% skimmed milk to block nonspecific immunoglobulin absorption to tissues. A heterologous hyperimmune serum from mouse infected with L. amazonensis was used as primary antibody. Slides were incubated for 18 h at 4 ◦ C in a humid chamber. After washing in PBS, the slides were incubated with biotinylated side antibody (DAKO), washed in PBS again, and the incubated with the streptavidin–peroxidase complex (DAKO) for 45 min each at 37 ◦ C. The reaction was developed with a diaminobenzidine (K3468, DAKO,) solution and hydrogen peroxide 10 (v/v). Finally, slides were dehydrated, cleared, counter-satined with Harris Hematoxylin, and mounted with coverlips. 2.6. Detection of FeLV antigen and FIV antibody To evaluate the impact of immunosuppressive retroviruses, all serum samples were tested for FeLV antigen (p27) and FIV antibody using a commercial rapid assay kit (SNAP® FIV Antibody/FeLV Antigen Combo Test: IDEXX Laboratories, Westbrook, ME, USA). 2.7. Real time polymerase chain reaction and sequencing To ensure that infection was caused by L. chagasi, EDTA anticoagulated whole blood samples were randomly obtained from positive cats, stored at −20 ◦ C and DNA extracted under standard protocols on a commercial platform (Corbett XTractor-Gene, Qiagen, Valencia, CA, USA). A housekeeping gene (18S rRNA) was used to determine DNA content and quality. The PCR test was based on IDEXX’s proprietary real-time PCR oligonucleotides (IDEXX Laboratories, Westbrook, ME, USA). Briefly, highly conserved metalloproteinase gp63 gene sequences were aligned and a region selected for 2 primers and a hydrolysis probe designed using commercial software (PrimerExpress Version 3.0, Applied Biosystems, Foster City, CA, USA). An additional 2 primers flanking the real-time PCR assay were designed for sequence verification purposes. Real-time PCR was run with standard primer and probe concentrations (ABI) using a commercially available mastermix (Roche LightCycler® 480 Probes Master, Roche Applied Science, Indianapolis, IN, USA). Real-time PCR was performed using default cycling conditions on a commercial instrument (Roche LC480 in the 384-well plate configuration, Roche Applied Science, Indianapolis IN, USA). Amplicons were submitted for sequencing according to standard protocols. 2.8. Statistical analysis Kappa test was used to verify the diagnostic test agreement. Differences were considered significant when the P value was <0.05. Statistical analysis was performed by a
Table 1 Results of the direct parasitological examination of the lymphoid organs (popliteal lymph nodes–PL, bone marrow–BM, spleen–S and liver–L); immunohistochemical reaction (IHQ) of skin lesions; ELISA and indirect immunofluerecence reaction (IFAT) for visceral leishmaniasis; and FIV specific antibodies detection in 27 cats naturally infected by L. chagasi living in an endemic area for visceral leishmaniasis in Brazil. Animal
Parasitological examination
ELISA
IFAT
IHQ
FIV
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27
+BM – +BM/ +S/ +L – – +PL/ +L – – – +BM/ +S +BM/ +S +PL/ +S +BM – – – +BM – +BM +PL – – – – – – –
− − − + + + + − + − + − − + + − − − − − + + + − + + +
− − − − + + + − − − − − + − − − − − − − − − − − + + −
− + + − − + − + − − − − − − + + − + − − − − − + + − −
− − − − − + − − − − − − − − − − − − − − − + + − + − −
commercially available software program (InStat® , GraphPad Software Inc., La Jolla, CA, USA).
3. Results 3.1. Study subjects A total of 55 cats presenting dermatological lesions, 17 males and 10 females with age average of 3.5 years old, were obtained from April, 2008 to November, 2009. From the total number of evaluated animals, 27/55 (49.1%) were positive for visceral leishmaniosis by at least one of the diagnostic methods applied.
3.2. Parasitological diagnosis Amastigote forms of L. sp. were identified in lymphoid organs from 10/55 (18.2%) infected cats (Table 1). The parasite was more frequently found in bone marrow (70.0%), followed by spleen (40.0%), lymph nodes (30.0%) and liver (20.0%). In five cats (50.0%) the parasite was identified in more than one organ. Three positive cats on parasitological test were also positive for ELISA, two for IFAT and one animal was considered positive by both serological techniques.
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3.3. ELISA and IFAT for L. sp. The calculated cut-off value used in the present study was 0.2765 for ELISA and 1:40 for IFAT. The seroprevalence of leishmaniosis found was 14/55 (25.4%) cats by ELISA and 6/55 (10.9%) cats by IFAT (Table 1). Levels of L. sp. antibodies in ELISA positive cats ranged from 0.278 to 1.631, with a mean of 0.710. Five IFAT positive cats had titers of 1:40 and one 1:80. From these six animals, five were also positive by ELISA. Amastigotes of L. sp were identified in lymphoid organs of 4/14 ELISA and 3/6 IFAT positive cats. A poor kappa overall agreement was found between serological tests and parasitological diagnosis ( = −0.006 and = 0.135, respectively) 3.4. Immunohistochemistry (IHC) The anti-Leishmania immunohistochemical assay allowed the identification of nine (16.4%) positive animals (Table 1). The parasite was identified on the tip of the pinna (55.5%), bridge of the nose (22.2%), chin area (11.1%) and around the eyes (11.1%). In five cats, although immunostaining of lesioned skin areas was positive, parasites were not identified in lymphoid organs and their serological tests resulted negative. 3.5. Detection of FeLV antigen and FIV antibody FIV specific antibodies were found in 6/55 cats (10.9%), from which 5/6 (83.3%) were Leishmania infected (Table 1). Amastigotes were evidenced in 3/5 (60.0%) cats with coinfection and all of them had anti-L. chagasi IgG antibodies, showing an association between FIV and L. chagasi specific antibodies. FeLV antigens were not detected in any cat.
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evident dermatological sign was the presence of alopecia observed in the cephalic region, particularly the pinna of 26 animals. Alopecia was also observed in the limbs, dorsal trunk, abdomen and tail. Ulcers with hemorrhagic crusts were observed in nine animals, eight of them simultaneously on the pinna and bridge of the nose. One animal initially presented alopecia and crusts only in the left pinna; after 60 days the cat tested positive and dermatological condition had evolved to deep ulcers with hemorrhagic crusts (Fig. 1). Erythema was found in eight cats on the pinna, abdomen, limbs, cervical area and bridge of the nose. One of these animals had a nummular erythematous alopecia measuring one centimeter in diameter on the bridge of the nose (Fig. 2), with positive fungal culture for Microsporum canis. The immunohistochemistry reaction of this area showed immunostaining for amastigote of L. sp. Hyperpigmentation, hyperkeratosis, lichenification and exudation on dorsal trunk in cephalic and abdominal region was observed in three animals (Fig. 2). One cat presented scaling, comedones and epidermal collarettes on dorsal trunk and pinna, characterizing a seborrheic dermatitis condition. A total of 19/27 (70.4%) Leishmania infected cats had systemic disorders along dermatological lesions, including lymphadenopathy (52.6%), dehydration (52.6%), weight loss (36.8%), diarrhea (21%), bilateral mucopurulent ocular discharge (15.8%), changes in the state of consciousness (15.8%), ulcerations in the oral cavity (10.5%), dyspnea (5.3%), bilateral purulent nasal discharge (5.3%) and corneal opacity (5.3%). Surprisingly, 8/27 (29.6%) cats had no clinical manifestation of systemic disease besides the dermatological lesions.
3.6. Real time polymerase chain reaction
4. Discussion
Real-time PCR was applied in samples from three cats. Average analytical sensitivity was ten molecules per reaction (reproducible in triplicate analysis). Analytical specificity was confirmed by sequencing a positive real-time PCR reaction. Analytical specificity was further confirmed by testing DNA extracted from 23 bacterial, protozoal and rickettsial isolates. Reproducibility (CV on CT values and on absolute quantified numbers), amplification efficiency, linearity, square values and signal to noise ratio parameters all passed the required criteria and thresholds. Nucleotide sequence obtained resulted in an identity level between 97% and 100% with XM 001463664.1, XM 001463663.1, XM 001463662.1, XM 001463661.1, AM 502228.1 sequences (geneBank), confirming L. chagasi diagnosis.
The outcome of 27/55 (49.1%) positive cats for Leishmania was significantly higher than previous studies in the same Brazilian endemic area, which have found 6/200 (6.5%), 8/200 (4.0%) and 3/52 (5.76%) positive cats (Rossi, 2007; Costa et al., 2010; Coelho et al., in press). The screening of cats with skin problems in our study caused a 7.5 to 12.3-fold increase in positivity, clearly showing the importance of dermatological signs for diagnosis of Leishmania. Our data was higher even when compared to highly endemic areas of other countries, such as 7/23 (30.4%) cats found in Portugal (Maia et al., 2008). In our study, amastigote forms of L. sp. was found in lymphoid organs of 10/55 (18.2%) cats, which is around 4.5fold higher than previously found in two cat surveys, with 8/200 (4.0%) (Costa et al., 2010) and 2/52 (3.84%) (Coelho et al., in press) positive samples. A reasonable explanation for this finding would be that once amastigotes have produced skin lesions, lymphoid tissues have probably been invaded as well. Nevertheless, skin problem screening has greatly increased Leishmania detection in cats. Divergent results were observed in amastigotes found in bone marrow (7/10; 70.0%) and lymph nodes (3/10; 30.0%) of cats with dermatological lesions when compared to bone marrow (3/8; 37.5%) and lymph nodes (7/8; 87.5%) in a general cat population previously studied (Costa et al., 2010).
3.7. Clinical signs Based on the topographic distribution of lesions, clinical signs included 15/27 (55.5%) cats with cephalic involvement, 6/27 (22.2%) with lesions on limbs, 6/27 (22.2%) on dorsal trunk, 3/27 (11.1%) on the tail and 3/27 (11.1%) on the abdomen. The most affected areas in the cephalic region were the pinna (60.0%), followed by generalized involvement (20.0%) and the bridge of the nose (20.0%). The most
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Fig. 1. Skin lesions in cats naturally infected by L. chagasi. (a) Alopecia on the external surface of the left pinna with erythema and crusts (b); the same animal showing, after 60 days, deep ulceration on the tip of the left pinna with hemorrhagic crusts (c); alopecia with active bleeding and hemorrhagic crusts on the base of the right pinna (d); the same animal, demonstrating that the lesions were bilateral and symmetric.
No reasonable cause has been found to explain the more likelihood of bone marrow for parasites to be found in the present study and therefore a higher sensitivity when compared to other lymphoid tissues. Interestingly, five cats had the diagnosis confirmed by just one of the four evaluated organs, which emphasizes the importance of simultaneous parasite searching for optimum sensitivity. Seroprevalence observed in the present study was significantly higher than previously found in randomly selected cats from the same area, which showed prevalence of 3% and 11.5% (Rossi, 2007; Costa et al., 2009). ELISA has been positive in 14/55 (25.5%) cats and IFAT in 6/55 (10.9%) cats, of which 5/6 were positive for both IFAT and ELISA. Levels of antibodies in ELISA positive cats ranged between 0.278 and 1.631, reaching up to 6-fold the cut-off in some cats as previously observed (Solano-Gallego et al., 2007).
Since serum samples of known negative cats were still reactive at a dilution of 1:20, the cut-off used to identify positive animals was 1:40, identical to previously obtained by others (Silva et al., 2008; Duarte et al., 2010). In the present study, IFAT positive cats all had low titers, varying between 1:40 (5/6) and 1:80 (1/6), differing from previous results of titers up to 1:320 in L. infantum naturally infected cats (Vita et al., 2005; Silva et al., 2008). The low agreements between the parasitological method and ELISA ( = −0.006), and the parasitological method and IFAT ( = 0.1356) were similar to those previously obtained (Rossi, 2007). Inconsistency when comparing results may be clearly explained since seven infected cats were only identified by ELISA, whereas only 4/14 ELISA positive cats had shown parasites in lymphoid organs. On the other hand, 3/6 IFAT positive cats
Fig. 2. Skin lesions in cats naturally infected by L. chagasi. (a) Alopecic, erythematous and circular area, of approximately one centimeter of diameter, in which amastigotes of the parasite were identified (b); cervical area of another cat exhibiting alopecia, erythema and exudation.
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had amastigotes detected on direct smears from lymphoid organs, in accordance of a recent study in which 91.3% of serologically reactive animals were parasitologically negative (Costa et al., 2010). Due to its low sensitivity, traditional histological diagnosis of leishmaniosis may be unsuccessful to detect the parasite. Immunohistochemical technique (IHC), on the other hand, is a highly sensitive and specific tool for the diagnosis of visceral leishmaniosis (Ferrer et al., 1988; Bourdoiseau et al., 1997). In the present study, the IHC for detection of L. sp. on the skin has increased the rate of positive results compared to the other performed methods. Amastigotes were easily observed by IHC in nine cats, five of them positive only by IHC, confirming previous descriptions in which the use of this exam may increase the possibility of finding parasites even when in low amounts in the evaluated tissues (Ferrer et al., 1988; Bourdoiseau et al., 1997). Skin parasitism may post an additional public health concern due to higher potential of infection to sand flies, since sand flies may be infected by L. infantum when feeding on a cat with visceral leishmaniosis, as recently demonstrated by xenodiagnosis (Maroli et al., 2007). In the present study, asymptomatic presentation was more frequent than symptomatic, which may lead public health professionals to underestimate the actual number of infected cats. This particular feature may be consequence of differences in immunological response of cats when compared to dogs. While dogs mainly present a humoral response, cats appear to have a higher degree of natural resistance due to a more cellular immune response (Solano-Gallego et al., 2007). In cases of immunosuppressive events, such as retrovirus infection, compromised cellular immunity may lead to the dissemination and visceralization of the parasite (Martín-Sánchez et al., 2007). Accordingly, 6/55 (10.9%) cats had anti-Feline Immunodeficiency Virus (FIV) antibodies, from which five (83.3%) had leishmaniosis, demonstrating an association between immunosuppression on FIV and Leishmania coinfection. These findings are in agreement with a previous study where a high L. infantum seropositivity rate in FIV-positive cats was reported (Pennisi et al., 2004). Since no FeLV antigens were detected in the present study, we were not able to determine whether FeLV infection is a potential risk factor for feline L. infantum infection as previously observed (Sherry et al., 2010). Alopecia, particularly observed on the pinnae, deep ulcerations and scaling were the most common dermatologic signs observed in this study. Some infected cats had only erythematous areas with alopecia or crusts without any nodules or ulcerated areas. These skin disorders may be associated with other dermatological conditions frequently affecting cats (Navarro et al., 2010). Consequently, leishmaniosis may be underestimated during clinical evaluation and differential diagnosis and clinicians should be aware of the concomitant presentation of dermatological problems, particularly when working in endemic areas. As previously observed, a total of 19/27 (70.4%) infected cats also showed systemic disorders such as lymphadenopathy, weight loss, mucopurulent ocular discharge, bilateral corneal opacity, purulent nasal discharge,
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changes in consciousness state, oral cavity ulcers, dyspnea and diarrhea (Poli et al., 2002; Vita et al., 2005). Since the aim of the study was strictly apply Real-time PCR to insure Leishmania species rather than a diagnostic approach, tests were performed only in three positive cats. L. chagasi strains were consistently identified by sequence analysis in all of them. Although two cases of L. amazonensis infection have been previously reported in domestic dogs living in the same area (Tolezano et al., 2007), two PCR positive cats were from the same shelter where 17 infected animals were found, and the third one was from other shelter were 10 infected animals were detected. However, further studies should be conducted in order to fully establish the Leishmania distribution in cat populations of endemic areas, as well as the cat role in the disease cycle. 5. Conclusion Based on the high association to dermatological lesions, leishmaniosis should be included as differential diagnosis of skin diseases of cats living in endemic areas. Moreover, immunohistochemistry should be always employed to increase diagnostic sensitivity. Finally, cats may represent a source of infection to sand flies as they have high skin parasitism. Acknowledgment We are grateful to the Fundac¸ão de Amparo à Pesquisa do Estado de São Paulo (FAPESP), project 2009/52812-3 for financial support. References Ayllon, T., Tesouro, M.A., Amusategui, I., Villaescusa, A., Rodriguez-Franco, F., Sainz, A., 2008. Serologic and Molecular evaluation of Leishmania infantum in cats from Central Spain. Ann. N. Y. Acad. Sci. 1149, 361–364. Baneth, G., 2006. Leishmaniasis. In Greene, C.E. (Ed.), Infectious diseases. Elsevier, Canadá, pp. 685–698. Bourdoiseau, G., Marchal, T., Magnol, J.P., 1997. Immunohistochemical detection of Leishmania infantum in formalin-fixed, paraffinembedded sections of canine skin and lymph nodes. J. Vet. Diag. Invest. 9, 439–440. Camargo, J.B., Troncarelli, M.Z., Ribeiro, M.G., Langoni, H., 2007. Canine visceral leishmaniosis: aspects of public health and control. Clín. Vet. 12, 86–92. Coelho, W.M.D., Pereira, V.B.R., Langoni, H., Bresciani, K.D.S. Molecular detection of Leishmania sp. in cats (Felis catus) from Andradina Municipality, São Paulo State, Brazil. Vet. Parasitol., in press. Costa, T.A.C., Rossi, C.N., Laurenti, M.D., Gomes, A.A.D., Vides, J.P., Sobrinho, L.S.V., Marcondes, M., 2010. Occurrence of leishmaniosis in cats of an endemic area for visceral leishmaniosis. Braz. J. Vet. Anim. Sci. 47, 213–217. Costa, T.A.C., Rossi, C.N., Laurenti, M.D., Gomes, A.A.D., Vides, J.P., Sobrinho, L.S.V., Costa, D.C., Marcondes, M., 2009. Use of enzyme-linked immunosorbent assay for serological diagnosis of Leishmaniasis in cats. In: Proceedings of the 34th WSAVA Congress , São Paulo, Brazil, pp. 93–94. Duarte, A., Castro, I., da Fonseca, I.M.P., Almeida, V., de Carvalho, L.M.M., Meireles, J., Fazendeiro, M.I., Tavares, L., Vaz, Y., 2010. Survey of infectious and parasitic diseases in stray cats at the Lisbon Metropolitan Area. Portugal. J. Feline Med. Surg. 12, 441–446. Ferrer, L., Rabanal, R., Fondevila, D., Ramos, J.A., Domingo, M., 1988. Skin lesions in canine leishmaniasis. J. Small Ann. Pract. 29, 381–388. Grevot, A., Jaussaud Hugues, P., Marty, P., Pratlong, F., Ozon, C., Haas, P., Breton, C., Bourdoiseau, G., 2005. Leishmaniosis due to Leishmania infantum in a FIV and FeLV positive cat with a squamous cell
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