African Journal of Health Sciences The Kenya Medical Research Institute (KEMRI) ISSN: 1022-9272 Vol. 13, Num. 1-2, 2006, pp. 7-21

African Journal of Health Sciences, Vol. 13, No. 1-2, Jan-June, 2006, pp. 7-21

Situational Analysis of Leishmaniases Research in Kenya

Willy Kiprotich Tonui

Kenya Medical Research Institute, P.O. Box 54840-00200, Nairobi, Kenya

Code Number: jh06003

SUMMARY

Leishmania spp are protozoan parasites of the Trypanosomatidae family that cause disease in humans and animals. In general, infections with these parasites can be divided into three main forms namely, cutaneous, mucocutaneous, and visceral leishmaniases. The disease is prevalent in many tropical and subtropical regions of the world, where it is transmitted via the bite of an infected sand fly. Leishmaniasis has been known to be endemic in parts of Kenya from as far back as early in the 20^th century. These endemic areas include Turkana, Baringo, Kitui, Machakos, Meru, West Pokot and Elgeyo Marakwet districts which have been reported to be endemic for kala-azar. Recent outbreaks of VL have been reported in the previously non-endemic districts of Wajir and Mandera in North Eastern Kenya between May 2000 and August 2001. The vector for VL in Kenya is Phlebotomus martini though other vectors including P. orientalis have been reported. Baringo district is the only foci reported where both VL and CL are known to occur in Kenya. The aetiological agents for CL which include L. major which has been reported in Baringo; L. tropica in Laikipia, Samburu, Isiolo, Nakuru and Nyandarua districts while L. aethiopica has been reported in the Mt Elgon area. In Kenya, P. duboscqi, P. guggisbergi have been shown to be the vectors of L. major and L. tropica, respectively, while P. pediffer, P. longipes and P. elgonensis have been implicated as vectors of L. aethiopica. Since 1980, the Kenya Medical Research Institute (KEMRI) has spearheaded research on leishmaniases research in Kenya focusing on various aspects including characterization of Leishmania species, biology, and ecology of sand fly vectors, development of biological strategies for sand fly control, identification of animal reservoirs, diagnosis, new treatment strategies, new chemotherapeutic agents, and vaccine-related studies. KEMRI, a founding partner of the Drugs for Neglected Disease Initiative (DNDi), whose overall aim is to address lack of new or improved drugs for neglected diseases (which include leishmaniases, malaria, trypanosomiasis and chagas disease) has made major contributions in leishmaniases research and control in Kenya and the eastern Africa region. [Afr J Health Sci. 2006; 13: 7-21]

Introduction

Leishmania spp are protozoan parasites of the Trypanosomatidae family that cause disease in humans and animals. In general, infections with these parasites can be divided into three main forms namely, cutaneous (CL), mucocutaneous (MCL) and visceral (VL) (or kala azar) leishmaniasis [1]. The disease is prevalent in many tropical and subtropical regions of the world, where it is transmitted via the bite of an infected sand fly. Although leishmaniasis is endemic in 88 countries, 90% of CL cases have been reported in Afghanistan Syria, Saudi Arabia, Brazil, and Peru. VL, also known as kala azar, is the most dangerous form of the disease and occurs in Bangladesh, Brazil, India, and Sudan. A new syndrome in which Leishmania co-infects patients with AIDS is particularly deadly and has been reported in 33 countries including Kenya [1, 2]. The impact of leishmaniasis on humans is enormous. One tenth of the world’s population is at risk and 2 million new cases are reported each year [1]. There are 22 species that cause disease in humans or animals [3] and specific parasite species are associated with specific types of syndrome (see Table 1). For example, VL is caused by L. donovani and L. chagasi [4] whereas CL is caused by different species in different areas of the world. For example, L. major causes CL in the Middle East and L. mexicana causes CL in Central America. In addition, lesions caused by L. braziliensis complex in the skin can spread to the mucous membranes resulting in a late complication known as MCL, or espundia [5].

Table 1: Spectrum of leishmaniases, aetiological agents and worldwide distribution

Epidemiology of leishmaniases in Kenya

Leishmaniasis has been known to be endemic in parts of Kenya from as far back as early in the 20^th century [6]. An outbreak of VL in the King’s African Rifles troops encamped north of Lake Turkana in southwest Ethiopia was reported in the 1940s [7]. Since then Turkana, Baringo, Kitui, Machakos, Meru, West Pokot and Elgeyo Marakwet districts have been considered to be endemic for kala-azar. Between 1950 and 1960 more than 1,052 cases of VL were reported from these districts alone [6].

A serious outbreak of VL was reported in Kitui district that began in 1952 with 303 cases being reported and peaked in 1953 with 2,142 cases. In the same year 157 cases were reported in the neighbouring Meru district [6]. Another outbreak of VL in Meru district was reported in 1966 with 1,500 cases [8]. Phlebotomus martini Parrot, 1936 was shown to be the vector of kala-azar during that epidemic [9, 10]. Further outbreaks of VL were reported in Machakos district in the 1970s, and again in Kitui district in the 1980s [11-13]. Baringo and the neighbouring districts such as West Pokot were first identified as leishmaniases foci in 1955 [14]. Since then, L. donovani transmitted by P. martini and L. major transmitted by P. duboscqi Neveu Lemaire, 1906 have been shown to exist in the region allopatrically, with rodents being the reservoir of CL (15-21). Other possible vectors that have been reported include P. celiae Minter, 1962, P. vansomeranae Heich, Guggisberg et Teesdale, 1956 and Sergentomyia garnhami [22] for VL [23, 24]. VL caused by L. donovani, is endemic in Baringo District, Kenya. The disease occurring in Baringo has a focal distribution in the dry, hot areas below 1500 metres and the infections may be characterized as follows: 1) asymptomatic 2) subclinical and self-limiting (not medically identifiable), and 3) clinically manifest disease (that is medically identifiable). Half of the reported VL patients are between 5 and 14 years of age and 66% of them are males. A human case of a mixed L. donovani and L. major infection has been reported in this dual focus of VL and CL [25]. Often associated with sporadic incidents of the disease rather than epidemics, as many as 305 cases of VL were reported in Baringo district alone in 1999 [26-29]. In the early 1990s, 3 Maasai children were diagnosed with VL acquired locally in Kajiado district in the Rift Valley of Kenya. L. donovani s.l. was isolated but a survey of other school children in the district did not reveal any more cases. P.martini was found to be common in the area while P. orientalis Parrot, 1936 another vector for VL though present was rare and therefore transmission was attributed to P. martini [30]. More recently, an outbreak of VL was reported in the previously non-endemic Wajir and Mandera districts of North Eastern Kenya where during a period from May 2000 to August 2001, 904 patients were diagnosed with VL, with the patients coming from an area that spanned Wajir and Mandera districts, southern Somalia and southeast Ethiopia. Unusual rainfall patterns, malnutrition and migration of a Leishmania-infected population seeking food and security are thought to have contributed to this outbreak [31].

Diffuse cutaneous leishmaniasis (DCL) was first described in Kenya in 1969 with 3 patients from the indigenous communities in Bungoma district, around the Mt. Elgon area diagnosed [32]. L. aethiopica has since been identified as the aetiological agent, rodents as the animal reservoirs and P. pedifer Lewis, Mutinga et Ashford, 1972 to be the vector of DCL in the Mt. Elgon region [14, 33, 34].

Cutaneous leishmaniasis caused by L. major, and transmitted by P. duboscqi was first reported in Kitui district in the 1990s [35].

L. major has also been isolated from an infected monkey (Cercopithecus aethiops) from Kiambu District, central Kenya [36]. However, no human cases have been reported from the district so far.

Meanwhile, active case detection and investigations of sand fly resting places conducted in the early 1990s, led to the identification of L. tropica CL in Central and Rift Valley provinces, mainly Laikipia, Samburu, Isiolo, Nakuru and Nyandarua districts [34]. Three sand fly species belonging to the subgenus Larroussius namely, P. pedifer, P. aculeatus Lewis, Minter et Ashford, 1974, P. guggisbergi Kirk et Lewis, 1952 and one Paraphlebotomus species (P. saevus Parrot et Martin, 1932) were found to be common in these areas [34,37-39]. P. guggisbergi has since been shown to be the vector for L. tropica in Kenya [40].

Contributions of the Kenya Medical Research Institute (KEMRI) to leishmaniases research and control in Kenya

Currently, disease control strategies rely on chemotherapy to alleviate the disease, on vector suppression, and personal protection to reduce transmission [1]. To date, there are no proven vaccines against any form of leishmaniasis [41]. Since 1980, the Kenya Medical Research Institute (KEMRI) has made major contributions to leishmaniases research and control focusing on various aspects including characterization of Leishmania species, biology, and ecology of sand fly vectors, development of biological strategies for sand fly control, identification of animal reservoirs, diagnosis, new treatment strategies, new chemotherapeutic agents, and vaccinerelated studies.

This has been achieved through collaborations with several agencies including the UN Development Program/World Bank/World Health Organization Special Program for Research and Training in Tropical Diseases (TDR), US Army Medical Research Unit-Kenya (USAMRU-K), the Ministry of Health’s Division of Vector Borne Diseases (DVBD), International Centre for Insect Physiology and Ecology (ICIPE), the Institute for Primate Research (IPR), Médecins Sans Frontières (MSF), local and international universities and colleges. As a result of this KEMRI scientists regularly participate in the Ministry of Health’s Disease Outbreak Management Unit, (DOMU), WHO and MSF to assist in the control of Kala azar outbreaks in Kenya and in the eastern African region. The Centre for Biotechnology Research and Development (CBRD) and the Centre for Clinical Research (CCR) of KEMRI have spearheaded leishmaniasis research. The Institute is a leader in sand fly biology research and it maintains the only sand fly colony in sub-Saharan Africa. The Centre for Clinical Research (CCR), designated a Good Clinical Practice (GCP) compliant facility, has also been involved in testing new antileishmanial drugs. KEMRI has an animal care facility that supports laboratory research in leishmaniasis.

The following sections summarize research activities on leishmaniases in Kenya in which KEMRI has been a major contributor.

Characterization of Leishmania species in Kenya

It is essential to know the identity of the parasite(s) in each focus, since this knowledge has implications for control and treatment of leishmaniases [42]. Therefore, isolates originating either from research, entomological and zoological activities should be identified and compared with international references stains. Cryobanks are also required for storage and for future availability. To address this need, a laboratory at CBRD of KEMRI was designated a regional reference centre for leishmaniasis taxonomy in Kenya. This facility maintains a cryobank of Leishmania isolates and is internationally referred to as Nairobi Leishmania Bank (NLB). At least 1800 Leishmania and other Leishmania-like isolates obtained from humans, other vertebrates, sand fly vectors and other arthropods from Kenya and other parts of Africa have been cryopreserved in this bank.

Through this bank it has been possible to compare the biological characteristics of the Leishmania parasites in Kenya and the neighbouring countries [43]. Zymodeme analysis of the Kenyan VL and CL isolates resulted in the identification of 11 subpopulations of the L. donovani species complex and six subpopulations of the L. tropica species complex endemic to different geographic areas of Kenya [37, 39]. A Leishmania-like isolate from the reptiles has since been identified as Sauroleishmania (RGER/KE/89/NLB-1236) and has been shown experimentally to cause L. major-like cutaneous lesions and visceral leishmaniasis in BALB/c mice [44].

Research on sand fly biology and control in Kenya

Control of phlebotomine sand flies remains a difficult problem throughout the world because of the insects' very highly specialized breeding sites. Use of insecticides remains the most effective method of sand fly control. However, resistance to insecticides has proved to be a major challenge. Use of bed nets or permethrin impregnated wall cloth, repellents and other personal protective measures have proved to be unreliable in Kenya [45]. With these shortcomings, there is need to identify new insecticides.

In general, control of leishmaniasis is hampered by the diversity of vectors, parasites, and reservoir hosts and the interventions must take into account these differences. It is therefore crucial to understand the biology of the leishmaniasis as well as that of the sand fly. The exploration of host-parasite interactions between species of Leishmania and respective vectors represents a wide and important field for basic and applied research as well. To address these needs, laboratory reared sand fly colonies have been established at KEMRI. These include P. martini and P. duboscqi Neveu-Lemaire (Diptera: Psychodidae) [17]. It is through studies done using P. duboscqi that L. major development within the sand fly was unravelled [46]. These studies have shown that it takes a period of 4-25 days, for the parasite to develop inside the sand fly and during the next blood meal the infective metacyclic stage enters the body of host through a sand fly bite [46, 47]. At least five developmental forms have also been recognized during development in the subgenus Leishmania, namely, procyclic promastigotes, nectomonad promastigotes, haptomonad promastigotes, paramastigotes, and metacyclic promastigotes [46].

Understanding of sand fly biology is also important in designing new control programmes in leishmaniases. It has also been shown that P. perniciosus can acquire and allow the development of L. infantum by feeding on L. infantum/HIV coinfected patients. Since this sand fly is an important vector of VL in southern Europe, a new natural anthroponotic cycle could be considered in the epidemiology of L. infantum/HIV coinfection [48]. This phenomenon is known as xenodiagnosis and this technique has been shown to be able to detect the presence of Leishmania parasites in a patient that other conventional diagnostic techniques failed (48). Therefore, the design of leishmaniasis control programs and the management of coinfected individuals should take these findings into account in southern Europe [48].

Investigations on vectors of leishmaniases in Kenya started only in the early 1950’s when the VL assumed importance as a result of a major disease epidemic outbreak [49]. Since then these investigations on vectors have resulted in identification of the main vectors for both VL and CL. P. martini Parrot has been confirmed as the vector for L. donovani the aetiological agent of VL in Kenya, southern Sudan, and Ethiopia [15, 21,50]. This sand fly species rests and breeds in termite hills, animal burrows, tree holes, and house walls [51, 52]. It feeds mainly on goats, rabbits, and humans, and to a lesser extent on dogs, cattle, and other hosts [52]. Another vector for L. donovani in Kenya is P. orientalis which occurs in Kajiado district of Kenya [30].

Among the cutaneous forms of leishmaniasis P. duboscqi Neveu-Lemaire (Diptera: Psychodidae) has been confirmed as the main vector of L. major in West Africa and the Rift Valley of Kenya [53, 54]. P. pediffer, P. longipes and P. elgonensis have been implicated as vectors of L. aethiopica occurring in the vicinity of Mount Elgon, Kenya [32, 34, 55, 56]. P. guggisbergi has been incriminated as a vector of L. tropica endemic in Samburu, Laikipia, Narok, Nakuru and Nyandarua districts of Kenya [37, 40].

Research on animal reservoirs of Leishmania in Kenya

Extensive research has been carried out in eastern Africa for animal reservoirs of both VL and CL. The domestic dog [54] has been the only domestic animal so far implicated as a possible reservoir for visceral leishmaniasis [58]. For cutaneous leishmaniasis caused by L. aethiopica, the rock (Denobrahyrax arboveus) and tree (Procavia johnstoni) hyraxes and the giant rat (Cricetomys sp) are the proven reservoirs of the disease [33], while several species of rodents have been demonstrated to harbor L. major which include Tatera robusta, Arvicanthis niloticus and Aethomys kaiseri [15, 18, 19]. The domestic sheep [58] and goats [59] which were initially suspected to be animal reservoirs for VL have since been proven experimentally to be non susceptible to L. donovani infection and therefore unlikely to be a synanthropic reservoir of this parasite [60, 61]

In Kenya, anthroponotic VL in Baringo District has shown that Leishmania deoxyribonucleic acid (DNA) is detectable in patients 10.5 months before diagnosis, up to 3 years after diagnosis and apparently successful treatment. Sub clinical cases have also been reported to have detectable circulating parasite DNA in their blood. These findings may indicate that sub clinical cases can be a reservoir and formerly treated VL patients can remain a reservoir for a long time. Xenodiagnosis may be important in such areas to determine whether sub clinical cases and former VL patients do have role in transmission of VL in Kenya [62].

Research on new diagnostic tests for leishmaniases

Sensitive, specific, reproducible, feasible and cheap diagnostic tests are needed if one wants to build an effective and efficient testtreatment strategy [63]. Since Leishman identified the parasite in 1901, the definitive diagnosis of VL has relied on demonstrating the organism by microscopy in smear or culture of aspirates or tissue [64,65]. However, a splenic-aspiration technique [66] which is used to acquire these tissues can hardly be recommended for rural district hospitals because of the risk of fatal haemorrhage, which subsists even with the recently introduced safer procedures [67]. Therefore, the search for a suitable alternative for splenic aspirates is thus still going on.

Over the past 20 years, several serological tests have been developed showing high sensitivity for visceral disease [68]. Enzyme linked immunosorbent assays (ELISA) for detecting leishmanial antibodies has also been established and successfully applied in clinical routine and seroepidemiological surveys in Kenya [26, 27, 69]. Of all the available serological tests, the direct agglutination test (DAT) has proved to be the most appropriate one for field use in Kenya [29, 63, 70, 71].

Other methods that have been tested include isoenzyme analysis [72, 73]; kDNA based techniques [74, 75]. In recent years polymerase chain reaction (PCR) techniques for leishmaniases have been developed and applied to visceral disease with contradictory results [76, 77]. Apart from the fact that PCR does not eliminate the need for tissue sampling, it is not exactly the type of appropriate technology needed in endemic areas.

Research on new treatment strategies and chemotherapeutic agents for leishmaniases

Neglected diseases such as leishmaniasis, trypanosomiasis, Chagas disease, and malaria have had a devastating impact on the world's poor. These treatable, infectious tropical diseases have been progressively marginalized by those in charge of making research programme decisions, both in the public and private sectors. Unfortunately, people suffering from these diseases do not constitute a market lucrative enough to attract investment in research and development for new drugs. KEMRI is a founding partner of an international effort, Drugs for Neglected Disease Initiative (DNDi), whose aim is to address lack of new or improved drugs for neglected diseases such as leishmaniasis. KEMRI’s Centre for Clinical Research (CCR) brings to DNDi a rich history of involvement in tropical diseases and expertise in clinical research for neglected diseases.

Over the last 25 years, scientists at CCR have been addressing the issue of treatment options for Kala-azar. The treatment of leishmaniasis, as currently conducted in Kenya with sodium stibogluconate, is unsatisfactory as it is expensive, resistance and relapses may occur [78]. However, it is through collaborative efforts of CCR scientists several new drugs or including AmBisome^® a safer and more effective lipid formulation of Amphotericin B (second line VL treatment); sitamaquine and aminosidine, alone or combined with sodium stibogluconate, in visceral leishmaniasis, compared to treatment by stibogluconate alone have been tested [79].

Worldwide, Leishmania parasites are increasingly becoming more and more resistant to antileishmanial drugs. The increment of in vitro antimony resistance due to intermittent drug exposure [80-83], the isolation of antimony-resistant Leishmania strains from patients with unresponsive CL [84,85], and recent reports, of numerous cases of VL among patients with the human immunodeficiency virus [86-89] indicate the need for new therapeutic agents for control and management of leishmaniases. Extensive studies of new drugs with antileishmanial activities, including natural products and synthetic compounds, have been undertaken worldwide [90], but problems associated with the currently available drugs remain unresolved. In recent years, there has been growing interest in alternative therapies and use of natural products, especially those derived from plants [91]. The use of plantbased medicine for healing and prophylaxis is as ancient and universal as medicine itself. Recent research undertaken in KEMRI has led to discovery of plant-based products with antileishmanial activities. For instance, studies have shown that leaf, root and stem extracts from Ajuga remota and Trichilla emetica have antileishmanial activity when tested against L. donovani in vitro and in vivo [92]. Some of the active compounds isolated from plants include iridoid glucosides, limonoids and terpenoids [92]. KEMRI has also initiated studies aimed at testing extracts/compounds from medicinal plants used widely by the local people for treating splenomegaly related ailments in leishmaniasis endemic areas of Kenya [92]. A long term goal of this research is to develop oral drugs for leishmaniases. These drugs have an advantage of reducing treatment-related socioeconomic difficulties (inadequate health care infrastructure and the long period of patient hospitalization) prevalent in the endemic areas.

Among the products tested for antileishmanial activity in the rodent models (BALB/c mice and Syrian hamsters) at KEMRI includes metal ion chelators namely (Pristane, EGTA, and HEEDTA). Results of these studies suggests that metal ions chelators have antileishmanial potential in vivo against L. donovani but their activity was low compared to that of Pentostam [93, 94].

Vaccine related research for leishmaniases in Kenya

Currently there is no vaccine available for use against any form of leishmaniases worldwide [41]. In brief, vaccines based on killed promastigotes with or without Mycobacterium bovis BCG have shown significant protection against L. braziliensis [95], L. mexicana or braziliensis, L. major [96] and L. donovani [97] in humans. In experimental animal models, several other vaccine preparations are being tested. Examples include: attenuated live parasites [98]; subunit vaccines delivered by live carriers such as BCG expressing the surface protease gp63 of L. major [99]; vaccinia virus expressing the glycoprotein gp46/M2/PSA-2 [100]; gp63 expressed in attenuated Salmonella typhimurium [101]; purified recombinant or native proteins formulated with an adjuvant such as the Leishmania-homologue of receptors for activated C kinase (LACK) plus IL-12 [103, 102], gp46/M2/PSA-2 or protein dp 72 plus Corynebacterium parvum [104], T cell epitopes plus Poloxamer 407 [104]; and vaccination with DNA encoding gp63 [105], LACK [106], or gp46/M2/PSA-2 [107]. In general, these vaccination protocols elicited partial protection against L. major.

Vaccine research at KEMRI is still at the rodent model stage of development in leishmaniases. So far live-attenuated L. major parasites [108], L. major culture-derived soluble exogenous antigens (SEAgs) [109], lipophosphoglycan (LPG) alone or LPG plus Mycobacterium bovis Bacille Calmette-Guerin (BCG) [110], and LPG plus salivary gland lysates [111] have been tested.

Transmission blocking vaccine studies in leishmaniasis have also been undertaken at KEMRI. These studies have shown that LPG is an excellent candidate as a transmission blocking vaccine against L. major infections [112,113]. In these studies, sand flies, which fed on BALB/c mice, immunized with L. major-derived LPG [114] or monoclonal antibodies raised against LPG [115] showed that parasite development was inhibited at the log phase (procyclic) of the parasite. There was also a marked reduction in the numbers of metacyclic promastigotes developing, leading to reduced transmission of L. major to naive BALB/c mice [113]. It has also been shown that P. duboscqi gut lysates and proteins present in L. major-derived LPG share two common proteins of molecular weights 105 kDa and 106 kDa [115]. The goal of these studies has been to develop a vaccine that can be utilized both to reduce transmissible infections within the sand fly and disease severity within the host. Some of the important findings relevant to vaccine research also include the determination of an inoculum dose required for effecting a visceral infection in rodents [116].

At the Institute for Primate Research (IPR) in Karen, Nairobi it has been shown that the vervet monkey (African Green monkey) is a good model for human cutaneous or visceral leishmaniases [117, 118]. Availability of a non-human primate model of leishmaniasis facilitates the study of different aspects of this disease and will accelerate the development of vaccines and new drugs for leishmaniases. Using this model the safety and immunopotency of the recombinant gp63 mixed with Baccille Calmette Guerin (BCG) vaccines [119], and also the adjuvant potential of two doses of IL-12 when used with a killed L. major vaccine [120] have been evaluated. It has also been demonstrated that high crossreactivity between L. donovani and L. major, and that L. donovani protects against L. major infections [121]. These findings have relevance for vaccine development in leishmaniases.

Conclusion

Although various aspects of the transmission and control of leishmaniases have been studied in Kenya, the impact of visceral leishmaniasis or kala azar is still enormous in Kenya. Control measures against sand flies and reservoir hosts need to be further evaluated in terms of their effect on disease incidence. Very few attempts have also been made to quantify local factors or to identify the causative mechanisms of epidemics in previously nonendemic or low endemic areas. Research to develop more effective drugs that meet the current standards of safety also remains a high priority. It is hoped that the entry of Drugs for Neglected Disease Initiative (DNDi), whose aim is to address lack of new or improved drugs for neglected diseases including leishmaniases will mobilize inter-country, international support and collaboration in terms of both finance and technical expertise towards achieving this goal. Diagnostic and vaccine-related studies are also still required to be pursued vigorously.

Many cases of leishmaniases still go unreported or undiagnosed hence the official statistics are currently not available for determining the actual number of cases in Kenya. Health education activities are still required to sensitize people living in endemic areas, medical and public health managers, and individuals at risk on leishmaniases. Decision makers should be sensitized on the importance of the leishmaniases as a public health problem so that they will support and defend control programmes. Notwithstanding these challenges, contributions from KEMRI and its partners towards leishmaniases control in Kenya and the eastern Africa region is commendable.

References

1. World Health Organization leishmaniasis control homepage: http://www. who.int/emc/diseases/leish/index.html.
2. Desjeux P. Leishmania / HIV coinfections. African Health 1995; 18(1):20- 2.
3. Marsella R; and Ruiz de Gopegui R. Leishmaniasis: a re-emerging zoonosis. International Journal of Dermatology 1998; 37: 801-814.
4. Hide G; and Tait A. The molecular epidemiology of parasites. Experientia 1991; 47:128-142.
5. Pirmez C. Immunopathology of American cutaneous leishmaniasis. Memorias do Instituto Oswaldo Cruz 1992; 5:105-109.
6. Fendall NR. The spread of kala-azar in Kenya. East African Medical Journal 1961; 38: 417-419.
7. Cole ACE; Cosgrove PC. and Robinson G. A preliminary report of an outbreak of kala-azar in a battalion of King’s African Rifles. Transactions of the Royal Society for Tropical Medicine and Hygiene 1942; 36: 25-34.
8. Wijers DJ. and Minter DM. Studies on the vector of kala-azar in Kenya. V. The outbreak in Meru district. Annals of Tropical Medicine and Parasitology 1966; 60(1): 11-21.
9. Heisch RB; Wijers DJ. and Minter DM. In pursuit of the vector of kala-azar in Kenya. BMJ, 1962; 5298: 1456-1458.
10. Wijers DJB. and Mwangi S. Studies on the vector of kala-azar in Kenya. VI. Environmental epidemiology in Meru district. Annals of Tropical Medicine and Parasitology 1966; 60(3): 373-391.
11. Ngoka JM. and Mutinga MJ. Visceral leishmaniasis in Kenya: The onset of an epidemic outbreak in the Machakos district of Kenya. East African Medical Journal 1978; 55(7): 328-331.
12. Ho M; Siongok TK; Lyerly WH. and Smith DH. Prevalence and disease spectrum in a new focus of visceral leishmaniasis in Kenya. Transactions of the Royal Society for Tropical Medicine and Hygiene 1982; 76(6): 741-746.
13. Wijers DJ. and Kiilu G. Studies on the vector of kala-azar in Kenya, VIII. The outbreak in Machakos District: epidemiological features and a possible way of control. Annals of Tropical Medicine and Parasitology 1984; 78(6): 597-604.
14. Mutinga MJ. Phlebotomus fauna in the cutaneous leishmaniasis focus of Mt. Elgon, Kenya. East African Medical Journal 1975; 52:307-7.
15. Heisch RB. The isolation of Leishmania from a ground squirrel in Kenya. East African Medical Journal 1957; 34:183.
16. Heisch RB; Grainger WE. and Harvey AE. The isolation of a Leishmania from gerbils in Kenya. Journal of Tropical Medicine and Hygiene 1959; 62(7): 158-159.
17. Beach R; Young DG. and Mutinga MJ. Phlebotomus (Phlebotomus) duboscqi from Kenya: a new record. Transactions of the Royal Society and Tropical Medicine and Hygiene 1982; 76: 707-708.
18. Githure JI; Beach RF. and Lightner LK. The isolation of Leishmania major from rodents in Baringo District, Kenya. Transactions of the Royal Society for Tropical Medicine and Hygiene 1984; 78: 283.
19. Githure JI; Schnur LF; Blancq SM. and Hendricks LD. Characterizations of Kenyan Leishmania spp. and identification of Mastomys natalensis, Tarterillus emini and Aethomys kaiseri as new hosts of L. major. Annals of Tropical Medicine and Parasitology 1986; 80: 501-507.
20. Muigai R; Githure JI; Gachihi GS; Were JB; Leuweenburg J. and Perkins PV. Cutaneous leishmaniasis caused by Leishmania major in Baringo District, Kenya. Transactions of the Royal Society for Tropical Medicine and Hygiene 1987; 81: 600-602.
21. Perkins PV; Githure JI; Mebrahtu Y; Kiilu G; Anjili C; Ngumbi PS; Nzovu J; Oster CN; Whitmire RE; Leeuwenburg J; Hendricks LD. and Koech DK. The isolation of Leishmania donovani from Phlebotomus martini collected in Baringo District, Kenya. Transactions of the Royal Society for Tropical Medicine and Hygiene 1988; 82: 695-700.
22. Heisch RB. Zoonoses as a study in ecology; with special reference to plague, relapsing fever, and leishmaniasis. British Medical Journal 1956; 32(4994):669-73.
23. Southgate BA. Studies in the epidemiology of East African leishmaniasis. 2. The human distribution and its determinants. Transactions of the Royal Society for Tropical Medicine and Hygiene 1964; 58: 377-390.
24. Kaddu JB. and Mutinga MJ. Leishmania in Kenyan Phlebotomine sand flies. II. Natural infection in the malphigian tubules of Sergentomyia garnhami and Sergentomyia antennatus. Insect Science and Applications 1984; 5: 239-243.
25. Schaefer KU; Kurtzhals JA; Sherwood JA; Githure JI; Kager PA. and Muller AS. Epidemiology and clinical manifestations of visceral and cutaneous leishmaniasis in Baringo District, Rift Valley, Kenya. A literature review. Tropical Geographical Medicine 1994; 46(3):129-33.
26. Jahn A. and Diesfeld HJ. Evaluation of a visually read ELISA for serodiagnosis and sero-epidemiological studies of kala-azar in the Baringo District, Kenya. Transactions of the Royal Society for Tropical Medicine and Hygiene 1983; 77(4):451-4.
27. Jahn A; Lelmett JM. and Diesfeld HJ. Seroepidemiological study on kala-azar in Baringo District, Kenya. American Journal of Tropical Medicine and Hygiene 1986; 89:91-104.
28. Mutero CM; Mutinga MJ; Ngindu AM; Kenya PR. and Amimo FA. Visceral leishmaniasis and malaria prevalence in West Pokot District, Kenya. East African Medical Journal 1992; 69: 3-8.
29. Mbati PA; Githure JI; Kagai JM; Kirigi G; Kibati F; Wasunna K. and Koech DK. Evaluation of a standardized direct agglutination test (DAT) for the diagnosis of visceral leishmaniasis in Kenya. Annals Tropical Medicine and Parasitology 1999; 93(7):703-10.
30. Johnson RN; Lawyer PG; Ngumbi PM; Mebrahtu YB; Mwanyumba JP; Mosonik NC; Makasa SJ; Githure JI. and Roberts CR. Phlebotomine sand fly (Diptera: Psychodidae) seasonal distribution and infection rates in a defined focus of Leishmania tropica. American Journal of Tropical Medicine and Hygiene 1999; 60(5):854-8.
31. Marlet MV; Sang DK; Ritmeijer K; Muga RO; Onsongo J. and Davidson RN. Emergence or re-emergence of visceral leishmaniasis in areas of Somalia, northeastern Kenya, and south-eastern Ethiopia in 2000-01. Transactions of the Royal Society for Tropical Medicine and Hygiene 2003; 97(5): 515-8.
32. Kungu A; Mutinga MJ. and Ngoka JM. Cutaneous leishmaniasis in Kenya. East African Medical Journal 1972; 48: 458 - 465.
33. Mutinga MJ. The animal reservoir of cutaneous leishmaniasis on Mt. Elgon, Kenya. East African Medical Journal 1975; 52: 142-151.
34. Sang DK; and Chance ML. Cutaneous leishmaniasis due to Leishmania aethiopica, on Mount Elgon, Kenya. Annals of Tropical Medicine and Parasitology 1993; 87(4): 349-357.
35. Mutinga MJ; Massamba NN; Basimike M; Kamau CC; Amimo FA; Onyido AE; Omogo DM; Kyai FM. and Wachira DW. Cutaneous leishmaniasis in Kenya: Sergentomyia garnhami (Diptera Psychodidae), a possible vector of Leishmania major in Kitui District: a new focus of the disease. East African Medical Journal 1994; 71(7): 424-428.
36. Binhazim AA; Githure JI; Muchemi GK. and Reid GD. Isolation of Leishmania major from a naturally infected vervet monkey (Cercopithecus aethiops) caught in Kiambu District, Kenya. Journal of Parasitology 1987; 73(6):1278-9.
37. Mebrahtu Y; Oster CN; Shatry AM; Hendricks LD; Githure JI; Rees PH; Perkins P. and Leeuwenburg J. Cutaneous leishmaniasis caused by Leishmania tropica in Kenya. Transactions of the Royal Society for Tropical Medicine and Hygiene 1987; 81: 923-924.
38. Mebrahtu Y; Lawyer P; Githure JI; Kagar P; Leeuwenburg J; Perkins P; Oster C. and Hendricks LD. Indigenous human cutaneous leishmaniasis caused by Leishmania tropica in Kenya. American Journal for Tropical Medicine and Hygiene 1988; 39:267-73.
39. Mebrahtu YB; Lawyer PG; Pamba H; Koech D; Perkins PV; Roberts CR; Were JB. and Hendricks LD. Biochemical characterization and zymodeme classification of Leishmania isolates from patients, vectors, and reservoir hosts in Kenya. American Journal of Tropical Medicine and Hygiene 1992; 47(6):852-92.
40. Lawyer PG; Mebrahtu YB; Ngumbi PM; Mwanyumba P; Mbugua J; Kiilu G; Kipkoech D; Nzovu J. and Anjili CO. Phlebotomus guggisbergi (Diptera: Psychodidae), a vector of Leishmania tropica in Kenya. American Journal for Tropical Medicine and Hygiene 1991; 44(3): 290-298.
41. Handman E. Leishmaniasis. Current status of vaccine development. Clinical Microbiology and Reviews 2001; 14(2): 229-243.
42. World Health Organization, Geneva (WHO) .Control of leishmaniasis. Report of a WHO Expert Committee. Technical Report Series 1990; 793.
43. Lewin S; Schonian G; El Tai N; Oskam L; Bastien P. and Presber W. Strain typing in Leishmania donovani by using sequenceconfirmed amplified region analysis. International Journal of Parasitology 2002; 32(10):1267-76.
44. Makau JK; Anjili CO; Dunton RF; Lugalia RM. and Ngeiywa MM. Cutaneous leishmaniasis in BALB/c mice caused by a Sauroleishmania species isolated from a plated lizard, Gerrhosaurus major (Squamata cordylidae). East African Medical Journal 1999; 76(9):501
45. Mutinga MJ; Renapurkar DM; Wachira DW; Basimike M. and Mutero CM. A bioassay to evaluate the efficacy of permethrin-impregnated screens used against phlebotomine sand flies (Diptera: Psychodidae) in Baringo District of Kenya. East African Medical Journal 1993; 70(3):168-70.
46. Lawyer PG; Githure JI; Anjili CO; Olobo JO; Koech DK. and Reid GD. Experimental transmission of Leishmania major to vervet monkeys (Cercopithecus aethiops) by bites of Phlebotomus duboscqi (Diptera: Psychodidae). Transactions of the Royal Society for Tropical Medicine and Hygiene 1990;84(2):229-32.
47. Killick-Kendrick R. The life-cycle of Leishmania in the sand fly with special reference to the form infective to the vertebrate host. Annales de parasitologie Humaine ET compare 1990; 65(1): 37-42.
48. Molina R; Lohse JM; Pulido F; Laguna F; Lopez-Velez R. and Alvar J. Infection of sand flies by humans coinfected with Leishmania infantum and human immunodeficiency virus. American Journal of Tropical Medicine and Hygiene 1999; 60(1):51-3.
49. Mutinga MJ. Phlebotomine sand flies (Diptera: Psychodidae): their importance and some aspects of control. East African Medical Journal 1996; 73(1):48-9.
50. Balkew M; Gebre-Michael T; Berhe N; Ali A. and Hailu A. Leishmaniasis in the middle course of the Ethiopian Rift Valley: II. Entomological observations. Ethiopian Medical Journal 2002; 40(3):271-82.
51. Mutinga MJ; Basimike M; Kamau CC. and Mutero CM. Epidemiology of leishmaniases in Kenya. Natural host preference of wild caught phlebotomine sand flies in Baringo District, Kenya. East African Medical Journal 1990; 67(5):319 - 27.
52. Ngumbi PM; Irungu LW; Robert LL; Gordon DM. and Githure JI. Abundances and nocturnal activities of phlebotomine sand flies (Diptera: Psychodidae) in termite hills and animal burrows in Baringo District, Kenya. African Journal Of Health Sciences, 1998; 5(1): 28-34.
53. Dedet JP; Derouin F; Hubert B; Shnur LF. and Chance ML. Isolation of L. major from Mastomys erythroleucus and Tatera gambiana in Senegal (West Africa). Annals of Tropical Medicine and Parasitology 1979; 73: 433-437.
54. Beach R; Kiilu G; Hendricks LD; Oster C. and Leeuwenburg, J. (Cutaneous leishmaniasis in Kenya: transmission of Leishmania major to man by the bite of naturally infected Phlebotomus duboscqi. Transactions of the Royal Society for Tropical Medicine and Hygiene, 1984; 78: 747-751.
55. Sang DK. and Sionkok TK. Cutaneous leishmaniasis and Phlebotomus longipes on Mount Elgon. East African Medical Journal 1983; 60: 826.
56. Mutinga MJ. and Odhiambo TR. Cutaneous leishmaniasis in Kenya. III. The breeding and resting site of Phlebotomus Pedifer (Diptera; Phlebotomine) in Mount Elgon focus, Kenya. Insect Science and Application 1986; 7: 175- 180.
57. Mutinga MJ; Ngoka JM; Schnur LF. and Chance ML. The isolation and identification of leishmanial parasites from domestic dogs in the Machakos District of Kenya, and the possible role of dogs as reservoirs of kala-azar in East Africa. Annals of Tropical Medicine and Parasitology 1980; 74(2):139-44.
58. Mutinga MJ; Kihara SM; Lohding A; Mutero CM; Ngatia TA. Karanu F. Leishmaniasis in Kenya: description of leishmaniasis of a domestic goat from Transmara, Narok District, Kenya. Tropical Medicine and Parasitology 1989; 40(2):91-6.
59. Williams AO; Mutinga J. and Rodgers M. Leishmaniasis in a domestic goat in Kenya. Molecular Cell Probes 1991; 5(5):319-25.
60. Anjili CO; Ngichabe CK; Mbati PA; Lugalia RM; Wamwayi HM. and Githure JI. Experimental infection of domestic sheep with culture-derived Leishmania donovani promastigotes. Veterinary Parasitology 1998; 31; 74(2-4):315-8.
61. Anjili CO; Olobo JO; Mbati PA; Robert L. and Githure JI. Experimental infection of domestic goats with Leishmania major through bites of infected Phlebotomus duboscqi and needle inoculation of culture-derived promastigotes. Veterinary Research Communication 1994; 18(4):301-5.
62. Schaefer KU; Kurtzhals JA; Gachihi GS; Muller AS. and Kager PA. A prospective sero-epidemiological study of visceral leishmaniasis in Baringo District, Rift Valley Province, Kenya. Transactions of the Royal Society for Tropical Medicine and Hygiene 1995; 89(5):471-5.
63. De Raadt P. Leishmaniasis and world health. In ecologie des Leishmanioses, Colloques Internationaux du NC.N.R.S. (Editors). Paris: Editions du C.N.R.S. 1977;. 239: 313-315.
64. Brycesson A. Leishmaniasis. In: Manson’s Tropical Diseases, 20^th Edition, Cook, G. C. (Editor). London: W.B. Saunders, 1996; 1213-1243.
65. Boelaert M; Criel B; Leeuwenburg J; Van Damme W; Le Ray D. and Van der Stuyft P. Visceral leishmaniasis control: a public health perspective. Transactions of the Royal Society for Tropical Medicine and Hygiene 2000; 94(5):465-71.
66. Kager PA; Rees PH; Manguyu FM; Bhatt KM. and Bhatt SM. Splenic aspiration: experience in Kenya. Tropical and Geographical Medicine 1983; 35: 125 -131.
67. Chulay JD. and Brycesson AD. Quantitation of amastigotes of Leishmania donovani in smears of splenic aspirates from patients with visceral leishmaniasis. American Journal Tropical Medicine and Hygiene 1983. 32(3): 475-479.
68. Kar K. Serodiagnosis of leishmaniasis. Critical Review in Microbiology 1995; 21: 123-152.
69. Okong'o-Odera EA; Kurtzhals JA; Hey AS. and Kharazmi A. Measurement of serum antibodies against native Leishmania gp63 distinguish between ongoing and previous L. donovani infection. APMIS 1993; 101(8):642-6.
70. Harith AE; Kolk AH; Kager PA; Leeuwenburg J; Faber FJ; Muigai R; Kiugu S. and Laarman JJ. Evaluation of a newly developed direct agglutination test (DAT) for serodiagnosis and seroepidemiological studies of visceral leishmaniasis: comparison with IFAT and ELISA. Transactions of the Royal Society for Tropical Medicine and Hygiene 1987; 81(4):603-6.
71. Boelaert M; El Safi S; Mousa H; Githure J; Mbati P; Gurubacharya V; Shrestha J; Jacquet D; De Muynck A; Le Ray D. and Van der Stuyft P. Multi-centre evaluation of repeatability and reproducibility of the direct agglutination test for visceral leishmaniasis. Tropical Medicine and International Health 1999; 4(1):31-7.
72. Mebrahtu Y; Lawyer P; Githure J; Were JB; Muigai R; Hendricks L; Leeuwenburg J; Koech D. and Roberts C. Visceral leishmaniasis unresponsive to pentostam caused by Leishmania tropica in Kenya. American Journal of Tropical Medicine and Hygiene 1989; 41(3):289-94.
73. Mebrahtu Y; Lawyer P; Githure J; Kager P; Leeuwenburg J; Perkins P; Oster C. and Hendricks LD. Indigenous human cutaneous leishmaniasis caused by Leishmania tropica in Kenya. American Journal of Tropical Medicine and Hygiene 1988; 39(3): 267-73.
74. van Eys GJ; Schoone GJ; Ligthart GS; Laarman JJ. and Terpstra WJ. Detection of Leishmania parasites by DNA in situ hybridization with non-radioactive probes. Parasitology Research 1987; 73(3):199-202.
75. Massamba NN. Mutinga MJ. Recombinant kinetoplast DNA (kDNA) probe for identifying Leishmania tropica. Acta Tropica 1992; 52(1):1-15.
76. Nuzum E; White F 3rd; Thakur C; Dietze R; Wages J; Grogl M. and Berman J. Diagnosis of symptomatic visceral leishmaniasis by use of the polymerase chain reaction on patient blood. Journal of Infectious Diseases 1995; 171(3):751-4.
77. Boelaert M and Dujardin JC. Diagnostic PCR with Leishmania donovani specificity. Tropical Medicine and International Health, 1999; 4: 789.
78. Nyakundi PM; Rashid JR; Wasunna KM; Were JB; Muigai R; Kirigi G. and Mbugua J. Problems in the treatment of kala-azar: case report. East African Medical Journal 1995; 72(6):406-8.
79. Chunge CN; Owate J; Pamba HO. and Donno L. Treatment of visceral leishmaniasis in Kenya by aminosidine alone or combined with sodium stibogluconate. Transactions of the Royal Society for Tropical Medicine and Hygiene 1990; 84(2):221-5.
80. Grogl M; Odula AMJ; Cordero LDC. and Kyle DE. Leishmania spp: development of pentostam-resistant clones in vitro by discontinuous drug exposure. Experimental Parasitology 1989; 69:78– 90.
81. United Nations Development Program/World Bank/World Health Organization. 1995. Tropical disease research. Progress 1975–94. Highlights 1993–94, p. 137. United Nations Development Program/World Bank/World Health Organization, Geneva.
82. Alexander JA; Stoskar R. and Russel DG. Leishmania species: models of intracellular parasitism. Journal of Cell Science 1999; 112: 2993-3002.
83. Bighetti E; Hiruma-Lima JS; Gracioso C A. and Brito AR. Anti- inflammatory and antinociceptive effects in rodents of the essential oil of Croton cajucara Benth. Journal of Pharmacy and Pharmacology 1999; 51:1447-1453.
84. Grogyl M; Thomson TN. and Franke ED. Drug resistance in leishmaniasis: its implications in systemic chemotherapy of cutaneous and mucocutaneous diseases. American Journal for Tropical Medicine and Hygiene 1992; 47(1): 117-126.
85. Berenguer J; Moreno S; Cercenado E; Bernaldo de Quiros JC; Garcia de la Fuente A. and Bouza E. Visceral leishmaniasis in patients infected with human immunodeficiency virus (HIV). Annals of International Medicine 1989; 111: 129–132.
86. Altes J; Salas A; Riera M; Udina M; Galmes A; Balanzat J; Ballesteros A; Buades J; Salva F. and Villalonga C. Visceral leishmaniasis: another HIVassociated opportunistic infections? Report of eight cases and review of the literature. AIDS 1991; 5:201–207.
87. Albrecht H; Stellbrink HJ; Gross G Berg G; Helmchen U. and Mensing H. Treatment of atypical leishmaniasis with interferon g resulting in progression of Kaposi’s sarcoma in an AIDS patient. Clinical Investigations 1994. 72:1041–1047.
88. Torre-Cisneros J. and Villanueva JL. Efficacy of liposomal amphotericin B in the treatment of visceral leishmaniasis in patients coinfected with the human immunodeficiency virus. Clinical Infectious Diseases 1995; 20: 191 (Letter.)
89. Alvar J; Cañavate C; Gutiérrez-Solar B; Jiménez M; Laguna F; López-Vélez, R; Molina R. and Moreno J. Leishmania and human immunodeficiency virus coinfection: the first 10 years. Clinical Microbiology and Reviews 1997; 10:298-319.
90. Croft SL. and Yardley V. Chemotherapy of leishmaniasis. Current Pharmaceutical Descriptions 2002; 8:319-342.
91. Rates SM. Plants as source of drugs. Toxicon 2001; 39:603-613.
92. Rono MK; Rukunga G; Ndiege I; Musau R; Were CL; Muthaura C; Ngumbi P. and Tonui WK. Bio-prospecting for antileishmanials from plants used for Kalaazar therapy by traditional healers in Baringo District of Kenya (Manuscript in preparation).
93. Mbati PA; Abok K; Orago AS; Anjili CO; Kagai JM; Githure JI. and Koech DK. Pristane (2,6,10,14-Tetramethylpentadecane) inhibits disease progression in Leishmania-infected Balb/c mice. African Journal of Health Sciences 1994; 1(4):157-159.
94. Mbati PA; Abok K; Anjili CO; Orago AS; Kagai JM; Githure JI. and Koech DK. Screening of metal ion chelators against Leishmania donovani-infected Syrian hamsters. African Journal of Health Sciences 1995; 2(1):223-227.
95. Armijos RX; Weigel MM; Aviles H; Maldonado R. and Racines J. Field trial of a vaccine against New World cutaneous leishmaniasis in an at-risk child population: Safety, immunogenicity, and efficacy during the first 12 months of follow-up. Journal of Infectious Diseases 1998; 177: 1352-1357.
96. Momeni AZ; Jalayer T; Emamjomeh M; Khamesipour A; Zicker F; Ghassemi RL; Dowlati Y; Sharifi I; Aminjavaheri M; Shafiel A; Alimohammadian MH; Hashemi-Fasharki R;. Nasseri K; Godal T; Smith PG. and Modabber F. A randomized, double blind, controlled trial of a killed L. major vaccine plus BCG against zoonotic cutaneous leishmaniasis in Iran. Vaccine 1999; 17: 466-472.
97. Khalil EA; ELHassan AM; Zijlstra EE;. Mukhtar MM; Ghalib HW; Musa B; Ibrahim ME; Kamil AA; Elsheikh M; Babiker A. and Modabber F. Autoclaved Leishmania major vaccine for prevention of visceral leishmaniasis: a randomized, double blind, BCG-controlled trial in Sudan. Lancet 2000; 356: 1565-1569.
98. Titus R; Gueiros-Filho F; de Freitas LA. and Beverley SM. Development of a safe live Leishmania vaccine line by gene replacement. Proceedings of the National Academy of Sciences USA1995; 92: 10267-10271.
99. Connell ND; Medina-Acosta E; McMaster WR; Bloom BR. and Russell DG. Effective immunization against cutaneous leishmaniasis with recombinant Baccille Calmette-Guerin expressing the Leishmania surface protease gp63. Proceedings of the National Academy of Sciences USA 1993; 90: 11473-11477.
100. McMahon-Pratt D., Rodriguez D; Rodriguez JR; Zhang Y; Manson K; Bergman C; Rivas L; Rodriguez JF; Lohman KL. and Ruddle NH. Recombinant vaccinia vaccine viruses expressing GP46/M-2 protects against Leishmania infection. Infection and Immunity 1993; 61: 3351-3359.
101. Xu D., McSolrley SJ; Chatfield SN; Dougan G. and Liew FY. Protection against Leishmania major infection in genetically susceptible BALB/c mice by gp63 delivered orally in attenuated Salmonella typhimurium (AroA^-AroD^-). Immunology 1995; 85: 1-7.
102. Mougneau E; Altare F; Wakil AE; Zheng S; Coppola T; Wang ZE; Waldmann R; Locksley RM. and Glaichenhaus N. Expression cloning of a protective Leishmania antigen. Science 1995; 268: 563-566.
103. Rachamin N. and. Jaffe CL. Pure protein from Leishmania donovani protects mice against both cutaneous and visceral leishmaniasis. Journal of Immunology 1993; 150: 2322-2331.
104. Spitzer N; Jardim A; Lippert D. and Olafson R. Long-term protection of mice against Leishmania major with a synthetic peptide vaccine. Vaccine 1999; 17: 1298-1300.
105. Xu D. and Liew FY. Genetic vaccination against leishmaniasis. Vaccine 1994; 12: 1534-1536.
106. Gurunathan S; Sacks DL; Brown DR; Reiner SL; Charest H; Glaichenhaus N. and Seder RA. Vaccination with DNA encoding the immunodominant LACK parasite antigen confers protective immunity to mice infected with Leishmania major. Journal of Experimental Medicine 1997; 186: 1137-1147.
107. Sjölander A; Baldwin TM; Curtis JM; Bengtsson KL. and Handman E. Vaccination with recombinant parasite surface antigen 2 from Leishmania major induces a Th1 type of immune response but does not protect against infection. Vaccine 1998; 16: 2077-2084.
108. Onyalo JA; Mwala DM; Anjili CO; Orago AS. and Tonui WK. Vaccinations with live-attenuated Leishmania major promastigotes and challenge infection with L. major in BALB/c mice. East African Medical Journal 2005; 82(4):193-7.
109. Tonui WK; Mejia JS; Hochberg L; Mbow ML; Ryan JR; Chan AS; Martin SK and Titus RG. Immunization with Leishmania major exogenous antigens protects susceptible BALB/c mice against challenge infection with L. major. Infection and Immunity 2004; 72(10):5654-61.
110. Tonui WK; Mpoke SS; Orago AS; Turco SJ; Mbati PA. and Mkoji GM. Leishmania donovani-derived lipophosphoglycan plus BCG induces a Th1 type immune response but does not protect Syrian golden hamsters (Mesocricetus auratus) and BALB/c mice against Leishmania donovani. Onderstepoort Journal of Veterinary Research 2003; 70(4):255-63.
111. Karanja R; Ingonga J; Mwangi M; Mwala D; Magambo J. and Tonui W. Immunization with a combination of Leishmania major Lipophosphoglycan (LPG) and Phlebotomus duboscqi salivary gland lysates (SGLs) abrogates protective effect of LPG against L. major in Balb/c mice. East African Medical Journal 2005; In Press
112. Tonui WK. Leishmania transmissionblocking vaccines: a review. East African Medical Journal 1999; 76(2):93-96.
113. Tonui WK; Mbati PA; Anjili CO; Orago AS; Turco SJ; Githure JI. and Koech DK. Transmission blocking vaccine studies in leishmaniasis. I: Lipophosphoglycan (LPG) is a promising transmission blocking vaccine candidate against cutaneous leishmaniasis East African Medical Journal 2001; 78(2):84-89.
114. Tonui WK; Mbati PA; Anjili CO; Orago AS; Turco SJ; Githure JI. and Koech DK. Transmission blocking vaccine studies in leishmaniasis: II. Effect of immunisation using Leishmania major derived 63 kilodalton glycoprotein, lipophosphoglycan, and whole parasite antigens on the course of L. major infection in BALB/c mice. East African Medical Journal 2001; 78(2):90-2.
115. Tonui WK; Ngumbi PM; Mpoke SS; Orago AS; Mbati PA; Turco SJ. and Mkoji GM. Leishmania major-Phlebotomus duboscqi interactions: inhibition of anti-LPG antibodies and characterisation of two proteins with shared epitopes. East African Medical Journal 2004; 81(2):97-103.
116. Mbati PA; Abok K; Orago AS; Anjili CO; Githure JI; Kagai JM. and Koech DK. Establishment of an appropriate inoculum dose of Leishmania donovani promastigotes required to establish a visceral infection in laboratory animal rodent models. African Journal for Health Sciences 1994; 1(4):165-168.
117. Githure JI; Reid GD; Binhazim AA; Anjili CO; Shatry AM. and Hendricks LD. Leishmania major: the suitability of East African nonhuman primates as animal models for cutaneous leishmaniasis. Experimental Parasitology 1987; 64(3):438-47.
118. Olobo JO; Gicheru MM. and Anjili CO. The African Green Monkey model for cutaneous and visceral leishmaniasis. Trends in Parasitology 2001; 17(12):588-92.
119. Olobo JO; Anjili CO; Gicheru MM; Mbati PA; Kariuki TM, Githure JI; Koech DK. and McMaster WR. Vaccination of vervet monkeys against cutaneous leishmaniosis using recombinant Leishmania 'major surface glycoprotein' (gp63). Veterinary Parasitology 1995; 60(3-4):199-212.
120. Gicheru MM; Olobo JO; Anjili CO; Orago AS; Modabber F. and Scott P. Vervet monkeys vaccinated with killed Leishmania major parasites and interleukin-12 develop a type 1 immune response but are not protected against challenge infection. Infection and Immunity 2001; 69(1):245-51.
121. Gicheru MM; Olobo JO. and Anjili CO. Heterologous protection by Leishmania donovani for Leishmania major infections in the vervet monkey model of the disease. Experimental Parasitology 1997; 85(2):109-16.

Copyright 2006 - African Forum for Health Sciences