Leishmania aethiopica: Experimental infections in non-human primates

Acta Tropica, 59 (1995) 243-250

243

© 1995 Elsevier Science B.V. All rights reserved 0001-706X/95/$09.50 ACTROP 00468

Leishmania aethiopica: Experimental infections in non-human primates Asrat Hailu a'*, Yohannes Negesseh and Isaac Abraham a alnstitute of Pathobiology (IPB), Addis Ababa University (AA U), P.O. Box 1176, Addis Ababa, Ethiopia bArmauer Hansen Research Institute (AHRI), P.O. Box 1005, Addis Ababa, Ethiopia (Accepted 7 March 1995) Six Cercopithecus aethiops monkeys, 4 Theropithecus gelada baboons and 2 Papio anubis baboons were infected using Leishmania aethiopica isolates originating either from localized (LCL) or diffuse (DCL) cutaneous leishmaniasis patients. The history of lesions in 4 C aethiops monkeys infected by LCL strains mimicked the process in human LCL patients. Infection of 2 C aethiops monkeys using a DCL strain resulted in localized, non-ulcerative, self-healing nodular lesions. Such lesions were also observed in 2 Z gelada baboons infected by LCL strains. Active lesions and healing in C. aethiops and T. gelada, after infection by LCL stains, were accompanied by positive DTH and immunity to challenge by LCL or DCL strains. Key words: Leishmania aethiopica; Non-human primate; Experimental cutaneous leishmaniasis; Animal model

Introduction

Leishmaniases in Ethiopia are caused by the species L. aethiopica, L. major and L. donovani sensu lato. L. major is considered to be the causative agent of CL in the lowlands of southwestern Ethiopia. L. aethiopica causes CL in the highlands and is widely distributed at altitudes between 1700-2700 meters (Ashford et al., 1977) . The disease is zoonotic, the parasite being maintained by two species of rock hyraxes i.e., P. capensis and H. brucei (Ashford et al., 1977). Transmission from rock hyraxes to man or from man to man occurs via two species of phlebotomines: P. longipes and P. pedifer (Lemma et al., 1969; Foster, 1972). CL due to L. aethiopica represents a unique variation of the Old World Leishmaniasis, since it exhibits a narrow hostparasite specificity between the rock hyraxes, the parasite and the two species of phlebotomines which are not known to serve as vectors of other leishmania. The parasite barely responds to sodium stibogluconate (Bray et al., 1973) and differs from all other leishmanias eg. by isoenzyme profile (Chance et al., 1978; Le Blancq *Corresponding author Abbreviations: Cutaneous leishmaniasis (CL); Localized Cutaneous Leishmaniasis (LCL); Diffuse Cutaneous Leishrnaniasis (DCL); Oriental Sore (0S); Delayed-type Hypersensitivity (DTH); Subcutaneous (sc); Intradermal (id); Post-infection (PI); Novy, MacNeal and Nicolle blood-agar slope medium (NNN); Deoxyribonucleic acid (DNA); Haematoxylin-Eosin (H & E).

SSDI 0001-706X(95)00085-2

244 et al., 1986) and by kinetoplastic DNA buoyancy and excreted factor serotype (Chance et al., 1978). The disease in humans presents as a spectrum ranging from localized self-healing to localized or diffuse non self-healing lesions. Natural L. aethiopica infections, other than in rock hyraxes, have been demonstrated in a few vertebrates e.g. Cricetomys sp. in Kenya (Mutinga, 1975). Childs et al. (1984) and more recently Akuffo et al. (1990) attempted unsuccessfully to infect white mice. Humber et al. (1989) used a variety of animals including white mice, but was only successful in infecting inbred hamster strains. Lack of animal models for this disease prompted us to carry out susceptibility experiments using three non-human primates namely C. aethiops, T. gelada and P. anubis. Preliminary results are presented and discussed.

Materials and methods

Fifteen green monkeys (Cercopithecus aethiops), 5 gelada baboons (Theropithecus gelada) and 3 anubis baboons (Papio anubis) were trapped from CL non-endemic areas. All animals and a baby born captive, were maintained in the animal house at the Institute of Pathobiology. The animals were kept in locally-made mobile individual cages and permitted once a day to move around and relax in an enclosed animal barn attached to the animal house. The regular diet included the staple grains (chickpeas, wheat, maize, green beans), cabbage, carrots, and bananas. Drinking water was provided twice a day with occasional vitamin and salt supplements. The animals were cared for and managed in the animal house looked after by the veterinary health department. All animals had been quarantined for natural infection with SIV, hemoparasites, intestinal parasites and leishmania for at least one year prior to inclusion in the experimental group. L. aethiopica infections were initiated in 6 C aethiops, 4 T. gelada , and 2 P. anubis. Two strains originating from LCL patients, strain # 1336/86 (AHRI ref #) and LES-20 (IPB ref #), and a strain originating from a DCL patient, LES-006 (IPB ref #) were used. These strains were characterized as L. aethiopica by isoenzyme electrophoresis technique (for 1336/86 and LES-006) or by restriction endonuclease analysis, southern blotting and using recombinant DNA probes (for LES-020) (van Eys et al., 1989). Stationary phase promastigotes grown on N N N medium were inoculated sc into the ear lobe, tip of nose or sides of the nose bridge at a dose of 5 x 106. All the infecting strains used for the experiment were recent isolates, passaged not more than five times in cultures. All animals, except two C. aethiops, were inoculated with LCL strains. Natural exposure/infection with leishmania was determined by detection of antileishmanial antibodies from sera collected serially over one year and by leishmanin skin test. Leishmanin was prepared locally. A single dose contained 5 × 10 6 L. aethiopica promastigotes in 0.1 ml of 0.5% phenosaline. The test was performed by id administration of the test antigen into the flexor surface of the shaved forearm. D T H reactions were read 48-72 hours after administration. Observation of the infected sites was made every third day to note emergence and appearance of lesions and subsequent changes. Lesional states were characterized as nodular, ulcerative or resolving on gross outcomes of infection. Lesion sizes were measured in millimetres of two diameters (length x width). Biopsies were obtained using a 3.0 mm biopsy

245 punch (Baker-Commins, USA), and fixed in 10% formol-saline. After routine processing of biopsy material, 5 ~tm sections were made and stained using H&E. Sections were examined to determine the extent of inflammatory infiltration and parasite density. Cellular infiltration was scored as absent (0), rare ( + / - ) , scanty (+), abundant (+ +) or numerous (+ + +). The density of parasites was scored as nondetectable (O),scanty ( + / - ) , abundant (+) or numerous (+ +). The presence and absence of granuloma and necrosis was determined. Parasite recovery was checked by making cultures of dermal scrapings or needle aspirates into N N N medium. Three of the six C. aethiops (AI, A2, and A3) which had spontaneously healed from primary infections with LCL strains and one T. galada (B~), were subjected to challenge infection with either LCL (in A1 and B~) or DCL (in A 2 and A3) strains. Challenge infections were performed parallel with new primary infections. The routes of administration, dose and culture stages (growth phases) of promastigotes were all as described for primary infection, only with changes in the sites of inoculations.

Results

Infections in all of the six C. aethiops and two of the four T. gelada produced recognizable lesions, but in none of the two P. anubis. Infection with LCL strains resulted in self-resolving ulcerative lesions in C. aethiops which progressed through the nodular, ulcerative and resolving phases. LCL strain infected T. gelada baboons produced non-ulcerative lesions. A DCL strain resulted in self-resolving, nonulcerative localized lesions in 2 C. aethiops. The pre-clinical period varied from 35-161 days in C. aethiops and 49-74 days in T. gelada, excluding infection with the DCL strain. The lifespan of lesions from emergence to healing varied from 42 to 126 days except in one C. aethiops (Az in Table 1) whose lesion lasted for up to 17 months, and DCL infections in two C. aethiops (As and A 7 in Table 1) where lesions persisted up to 10-13 months after emergence. In A1 (Table 1), where the inoculum was administered in three divided doses, lesions were multiple and progressed through an isophasic development to ulceration and healing. Maximal lesion sizes were attained within the first third of the lifespan of active lesions. The maximum lesion size was observed among C. aethiops infected with a DCL strain (Table 1). Scars were formed after complete healing of ulcerating lesions and were typical of CL scars in humans, i.e., hyperpigmented and depressed with elevated margins. Healing in Az started at around days 221 PI (71 days after emergence) and was not complete until 660 days PI (Table 1). The healing stage persisted for 439 days, bringing lesion lifespan to 510 days when completely healed. A2 was later found to be pregnant when it gave birth. The healing stage persisted until 150 days after parturition and was completely healed by the end. All challenge infections were resisted. In A2 however, a transient nodule resulted 42 days post-challenge and was cleared within 32 days. DTH responses, monitored with sequential testing through the various stages of the disease, revealed that A1, A2, A3 and Ba (all infected with LCL strains) converted positive at early stages during the nodular state of lesions. C1, C2, Bz, B3 and B5 (all infected with LCL strains) and A5 and A7 (infected with a DCL strain) were all skin test negative throughout. Concurrent repeated administration of leishmanin in

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247 three control C aethiops monkeys (A,o, A~I, A~2) did not result in positive DTH conversions or immunity to primary infection. A~o, A~I, A~z were later successfully infected with an LCL strain (Strain # LES - 020). Inflammatory mononuclear cell infiltration was the basic feature of all lesions (Table 2). The infiltrate was mainly in the deeper dermis in nodular lesions of As, Ba, and Bs. Parasites were scanty or non-detectable with regressing nodules, late ulcers or scars eg. in A2 and A5 (Table 2). In B2, a dense infiltrate of histiocytes heavily packed with amastigotes was noted. A predominantly lymphohistiocytic infiltration was more clearly shown in the ulcerative lesions of A1, and A3 as well as in the nodules of A2, B 2 and B5 (Table 2). In the deep dermis of a regressing nodule from A 5, there was a moderate lymphohistiocytic infiltrate (Table 2). The upper and middle dermis were normal and remnants of dead L.d. bodies were visible in the deep dermis. Subsequent biopsies from disappearing lesions revealed chronic perivascular inflammatory infiltrate. Tuberculoid tissue responses and hence granuloma was not a typical finding. A biopsy obtained from a transient nodule, following challenge by a DCL stain in A2, revealed a tuberculoid granulomatous reaction composed of giant cells, epithelioid cells and lymphohistiocytic infiltrate. Attempts to recover parasites on NNN medium were successful with early active lesions (Table 2).

Discussion

This paper demonstrates that C aethiops is susceptible to infection with L. aethiopica. The susceptibility of C aethiops to L. major infection has earlier been reported (Githure et al., 1987; Lawyer et al., 1990). Infection of C aethiops by LCL strains had resulted in OS-like lesions which in the end self-resolved, accompanied with positive DTH and immunity to challenge infections. In T. gelada, lesions failed to ulcerate and the histology revealed a picture reminiscent of early OS nodules. The inflammatory cell infiltrate in C aethiops lesions is consistent with human CL Lesions of L. aethiopica infections. The incubation period, lifespan of lesions and duration of healing were variable as is also the case in humans. Lemma et al. (1969) reported that onset to healing of lesions in humans occurs in less than 2 years, and ulceration within 3 - 5 months after appearance of nodules. Emergence of lesions in our experimental animals was preceded by an incubation period of 1 - 5.5 months. Healing occurred in as little as 42 days or remained active for over a year. We have attributed the long persistence of lesions in A2 to immunodepression associated with pregnancy. The single nonulcerative, non-disseminative, self-resolving nodules observed in two C aethiops after infection with a DCL strain, is inconsistent with the picture in human DCL or LCL. It is unclear whether the lesions were of the early OS-type or primordial DCL lesions. DTH was negative, cutaneous metastases to local or distant sites was abortive (local spread of lesion on A5 to a bilaterally symmetrical site on the sides of the nose bridge was short-lived). These DCL-like lesions have disappeared and it is yet to be observed if secondary lesions would emerge. Bryceson (1969) provided the history of early DCL lesions, as obtained from recall memories of patients, and described the variability in the rates of spread to local or distant sites. Twenty years was the example given for late metastases.

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That a DCL strain resulted in similar DCL-like lesions in the two monkeys A5 and AT and also that two LCL stains resulted in LCL-like lesions in four monkeys A1, A 2, A 3 and m4, suggests that strain differences are responsible for the clinically distinct entities in L. aethiopica infections. Opposing views are expressed as to whether DCL and LCL are outcomes related to parasite or host factors. A longterm follow-up of our experiment will provide further data on infection with DCL strains. C. aethiops is widely distributed in East Africa, including CL endemic areas in Ethiopia. The possibility that it serves as a secondary host of the disease needs investigation. The species is here shown to be susceptible to experimental infection by L. aethiopica. Further experimental studies are warranted to explore its potential as a primate model of CL due to L. aethiopica.

Acknowledgements We are grateful to Messrs Kassa Bemnet and Kiros Ayenew for processing the biopsies. We thank Dr. Tivadar L. Miko for his help in the examination of histological specimens. We thank Dr. D.A. Evans of the School of Hygiene and Tropical Medicine (London), Dr. M. Gramiccia of the Istituto Superiore di Sanita (Rome) and Dr. G.J.J.M. van Eys of the Royal Tropical Institute (Amsterdam) for typing our strains. We also thank very much Drs. Dominique Frommel and Beyene Petros for their continued moral support and interest in the project. Messrs. Tesfaye Getachew and Mulugeta Gichile are also thanked for their participation in animal trapping and maintenance. We thank Dr. Sally Cowley for the linguistic corrections. The study was carried at the Institute of Pathobiology (IPB), Addis Ababa University (AAU) and the Armauer Hansen Research Institute (AHRI) Addis Ababa, Ethiopia. AHRI is supported by the Norwegian Universities Committee for Development Research and Education (NUFU).

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