Brugia malayi

Brugia malayi (Brug, 1928) Buckley, 1960

(Figures 4-50 through 4-51)

ETYMOLOGY: Brugia for Dr. Brug and malayi for the area in which the parasite was initially isolated.

SYNONYMS:Filariamalayi Brug, 1927; Microfilariamalayi (Brug, 1927) Faust, 1929; Filariabancrofti Cobbold, 1877; Wuchereriamalayi (Brug, 1927) Rao and Maplesonte, 1940.

HISTORY: Brugiamalayi was first described as Filariamalayi by Dr. SL Brug on the basis of the morphology of microfilariae that were found in people in Indonesia by a Dr. A Lichtenstein (Brug, 1928). Dr. Lichtenstein had noted that unlike the human parasite Wuchereriabancrofti in Indonesia, the microfilariae of this parasite did not increase in the peripheral circulation nocturnally and were not infective to culicine mosquitoes (Lichtenstein, 1927). Rao and Maplestone (1940) were the first to describe the adults of this species. Buckley (1960) erected a new genus, Brugia, on the basis of specimens of Brugiamalayi from monkeys, Brugiapahangi from cats, dogs, and monkeys in Malaysia, and Brugiapatei from cats and dogs in East Africa.

GEOGRAPHIC LOCATION: The location of Brugiamalayi in Asia is illustrated by Denham & McGreevy (1977), but this illustration is based mainly on reports in people. Basically this region includes India, Sri Lanka, Sumatra, Java, Borneo, Malaysia, the Philippines, southern Thailand and northern Vietnam, South Korea, and coastal China. Cats only become infected with Brugiamalayi in areas where this species is present in a subperiodic form, i.e., microfilariae are present in the blood throughout all hours of the day. This subperiodic form is present in Malaysia, Thailand, Vietnam, Borneo, Java, and the Philippines. In these areas, several studies have identified circulating microfilariae in cats. Chang et al. (1992) found 10.5% of 191 domestic cats positive for Brugiamalayi in 12 Malay villages in Sarawak, Malaysia, and Mak et al (1980) found infected cats in peninsular Malaysia. Dondero and Menon (1972) found 2 of 9 cats in Perak Malaysia with circulating microfilariae. Palmieri (1985)in South Kalamantan Indonesia found 4 of 325 cats positive; Partono et al (1977) found 13 of 51 cats positive). Phatana et al. (1987) found a cat positive for Brugiamalayi in southern Thailand.

LOCATION IN HOST: The adult worms are found in the lymphatic vessels as also are larval stages (Ahmed, 1966). Ewert (1971) showed that if infective-stage larvae were inoculated into the foot of the cat, the worms developed within the popliteal lymph node of that same leg, only very rarely were worms recovered from other sites.

PARASITE IDENTIFICATION: The adult males of Brugiapahangi are 13 to 23 mm long; the adult females of Brugiapahangi are 43 to 55 mm long (Buckley & Edeson, 1956). The adults of Brugiapahangi are most easily recognized by the male spicules; those of those of Brugiamalayi are the longest, those of Brugiapatei are intermediate in length, and the spicules of Brugiapahangi are the shortest of the Brugia spp. in cats. The left spicule of Brugiamalayi is 390 µm; the right spicule is 125 µm long. There is probably no good way to distinguish the females of these Brugia species.

Diagnosis of infection is made by finding the microfilariae in the blood using a Knott’s technique or by direct smear (Figs. 4-50 & 4-51). The microfilaria of Brugiamalayi is 177 µm to 230 µm long when examined in a thick blood film. In humans, the microfilariae of the subperiodic strain differs from that of the periodic form in that the microfilaria of the subperiodic form tends to lose its sheath in the process of dying on slides. The numbers of circulating microfilariae per milliliter of blood appears to remain lower than that of the microfilariae of cats infected with Brugia pahangi. Burren (1972) found in experimentally infected cats that there was around a maximum of <1 to only 3 microfilaria per μl of blood. It is difficult to distinguish the microfilaria of Brugiamalayi from that of Brugiapahangi. The distinction between the two species is made by the examination of the innenkorper of Giemsa stained microfilariae, the innenkorper in the microfilaria of Brugiamalayi is shorter than that in Brugiapahangi (Sivanandam and Fredericks, 1966). Also, when the microfilariae are stained using the acid phosphatase histochemical method, the microfilaria of Brugiamalayi are red mainly at the excretory and anal pores while those of Brugiapahangi tend to be red throughout their length (Redington et al., 1975). An antigen detection ELISA and counter-immunoelectrophoresis have been used to detect infections of Brugiapahangi in cats (Au et al., 1981; Kumar et al., 1991), but these tests are not commercially available. Pasomsitti et al (1983) used an indirecte fluorescece technique to examine antibody levels in cats infected with Brugiamalayi, Brugiapahangi, or Dirofilariarepens.

LIFE CYCLE: Ewert (1971) examined the development of Brugiamalayi in experimentally infected cats. If larvae were inoculated into the hind leg, the majority of adult worms were recovered from lymph nodes associated with that same leg. In one cat, examined 16 weeks after infection, worms were recovered from soakings of the skin of the leg and a single female worm was recovered from soakings of the scrotum and testes. Ewert (1976) found no difference as to the susceptibility of male and female cats to infection; also , out of 59 female and 41 male cats that were experiemntally infected, microfilariae appeared in the blood of about 90% (92% in the females and 88% in the males) of all these cats.

In cats infected with third-stage infective larvae from mosquitoes, the prepatent period ranged from 80 to 96 days (Edeson and Buckley, 1959; Edeson and Wharton, 1957; Wharton et al., 1958). The inoculated third-stage larvae reach the regional lymph glands and vessels within 16 hours, and typically, as also shown by Ewert (1971), the site of adult development was typically realated to the site of inoculation. The mole from third stage to fourth stage occurred about 10 days after the cats were infected, and the molt to the adult stage takes place 35 to 40 days after infection. Ewert and Bosworth (1975) described the results of experimentally exposing cats to a second inoculation at various times after the primary inoculation. The first infection did not consistently result in lowered numbers of developing worms in the same leg, but the worms that did develop were typically further from the inoculation site on the foot. Also, the primary infection had no apparent effect on the early migration and development of worms when the second infection was in the other hind leg.

The development in mosquitoes was described by Feng (1936). The first molt took place four days after the mosquito fed on the infected blood, and the second molt took place 2 days after the first molt. The infective larvae were 1.3 mm long..

CLINICAL PRESENTATION AND PATHOGENESIS: There have been no reports detailing the clinical presentation of naturally acquired Brugiamalayi infection in cats. Ewert et al (1972) examined the lymphographic changes in young cats that had been experimentally infected in the hind feet. After infection, lymphatic vessels in the infected limb become occluded by developing worms, and collateral lymphatics develop to maintain drainage. Eventually infested lymphatic vessels become obliterated (Folse et al., 1981). In affected limbs, the collagen content increases (Dresden & Ewert, 1984), and in experimentally infected cats thrombi within the lymph vessels slowly turn into fibrous tissue that occludes the flow of lymph (Fader & Ewert, 1986). Dependent limb edema develops in cats with experimental infections that can be grossly visible (Folse & Ewert, 1988).

In an attempt to explain why some humans infected with Brugiamalayi develop elephantiasis while the majority of those infected do not, work was performed to examine the effects of secondary bacterial infections on the course of infection of Brugia malayi in cats. When cats infected in one hind leg were challenged with streptococci in both hind legs, serious complications developed only in the previously worm-infected leg (Bosworth & Ewert, 1975). When similar conditions were extended to over a year, an elephantoid condition developed in the legs of 5 of 6 cats that had been repeatedly exposed to Brugia malayi and streptococcus (Ewert et al, 1980).

TREATMENT: In human infections with Brugiamalayi, treatment has been aimed at giving both diethylcarbamazine (DEC) and ivermectin at levels that are low enough to prevent the circulation of microfilariae for a year without the induction of major side effects amongst infected individuals. In other cases, DEC has been added to salt in endemic areas (Panicker et al., 1997). There have been no reported attempts to either treat or control infections in naturally infected cats.

Ewert and Emerson (1979) reported on the effects of DEC on fourth-stage and adult Brugia malayi in experimentally infected cats. When the cats were treated 20 days after infection with 100 mg/kg of DEC, no worms were present in the treated cats. When the cats were treated with 50, 25, or 10 mg/kg of DEC at 20 days after infection, worms were present in all cats at necropsy, although less than in the control groups. Treatment with 1 mg/kg had no effect on the number of worms developing. When the cats were treated 8 weeks after infection, 2 of the 5 treated cats harbored 1 or 2 adult worms, while the mean number of worms present in the control cats was 23 adults. The results of treatment at 1, 10, 25, and 50 mg/kg were similar to those reported for larvae (Hillman et al., 1983).

EPIZOOTIOLOGY: Relative to the cat, the most important aspect of the epizootiology of this disease is that cats are only infected naturally in those areas of the world where people are infected with the subperiodic form of the disease. Thus, throughout most of the range of this parasite, cats are not reservoirs of the infection. The cat can be experimentally infected with the nocturnally periodic form of the parasite, and in the experimentally infected cat, the periodic form becomes subperiodic (Denham and McGrreevy, 1977; Laing, 1961). This means that cats entering an area where the nocturnal form of the disease is prevalent may be at some risk of becoming infected.

HAZARDS TO OTHER ANIMALS: Various primates and species of wild felid are considered to be susceptible to infection with Brugia malayi. Thus, if cats are held in places where they may be bitten by mosquitoes, the parasite could be transmitted from an infected cat to one of these other hosts.

HAZARD TO HUMANS:Brugiamalayi is a significant enough human pathogen to warrant separate sections in most human parasitology texts. It is very much unclear what role the cat plays in the infection of people with Brugiamalayi. It is assumed that they serve as reservoirs in Malaysia and nearby countries, but it is truly not clear whether the cats or the human beings are the actual reservoirs of the observed infections.

CONTROL/PREVENTION: Oral ivermectin administered to four leaf monkeys, Prebytiscristata, at doses of 200 μg/kg to 300 μg/kg at the same time as the subcutaneous inoculation of infective larvae failed to prevent the development of patent infections in these animals (Mak et al., 1987). On the other hand, diethylcarbamazine appears to be highly effective in preventing the infection of cats with this parasite (Ewert & Emerson, 1975). In cats that were experimentally infected with 50 larvae, the treatment of the cats the first week after infection with 10 mg of diethylcarbamazine per kg body weight caused only 1 of 22 cats to have any detectable living larvae at necropsy two weeks after infection. When cats were administered 5 mg diethylcarbamazine per kilogram body weight, a single living larva was recovered from each of 2 of the 5 treated cats. It would thus seem warranted to begin cats on daily diethylcarbamazine at 6 mg per kilogram body weight (e.g., Filaribits) before taking them into a country where this parasite is endemic.


Ahmed SS. 1966. Location of developing and adult worms of Brugia sp. in naturally and experimentally infected animals. J Trop Med Hyg 69:291-293.

Bosworth W, Ewert A. 1975. The effect of streptococcus on the persistence of Brugiamalayi and on the production of elephantiasis in cats. Int J Parasitol 5:383-389 9maybe 583-589).

Buckley JJC, Edeson JFB. 1956. On the adult morphology of Wuchereria sp (malayi?) from a monkey (Macacairus) and from cats in Malaya, and on Wuchereriapahangi n. sp. from a dog and a cat. J Helm 30:1-20.

Burren CH. 1972. The behaviour of Brugiamalayi microfilariae in experimentally infected domestic cats. Ann Trop Med Parasitol 66:235-242.

Chang Ms, Ho BC, Hardin S., Dorasisingam P. 1992. Filariasis in Kota Samarahan District Sarawak, East-Malaysia. Trop Biomed 9:39-46.

Dondero TJ, Menon VVV. 1972. Clinical epidemiology of filariasis due to Brugiamalayi on a rubber estate in West Malaysia. SE Asian J Trop Med Pub Hlth 3:355-365.

Dresden MH, Ewert A. 1984. Collagen metabolism in experimental filariasis. J Parasitol 70:208-212.

Edeson JFB. 1959. Studies on filariasis in Malaya: the periodicity of the microfilariae of Brugiamalayi and B. pahangi in animals. Ann Trop Med Parasitol 53:381-387.

Edeson JFB, Buckley. 1959. Studies on filariasis in Malaya: on the migration and rate of growth of Wuchereriamalayi in experimentally infected cats. Ann Trop Med Parasitol 53:113-119.

Edeson JFB, Wharton RH. 1957. The transmission of Wuchereriamalayi from man to the domestic cat. Trans Roy Soc Trop Med Hyg 51:366-370.

Edeson JFB, Wharton RH. 1958. The experimental transmission of Wuchereriamalayi from man to various animals in Malaya Trans Roy Soc Trop Med Hyg 52:25-45.

Edeson JFB, Wilson T. 1964. The epidemiology of filariasis due to Wuchereriabancrofti and Brugiamalayi. Ann Rev Entomol 9:245-268.

Ewert A. 1971. Distribution of developing and mature Brugiamalayi in cats at various times after a single inoculation. J Parasitol 57::1039-1042.

Ewert A. 1976. The comparative susceptibility of male and female and of mature and immature cats to infection with subperiodic Brugiamalayi. Rev Biol Trop 24:262-266.

Ewert A, Balderach R, Elbiharia S. 1972. Lymphographic changes in regional lymphatics of cats infected with Brugiamalayi. Am J Trop Med Hyg 21:407-414.

Ewert A, Bosworth W. 1975. Distribution and development of Brugiamalayi in reinfected cats. J Parasitol 61:610-614.

Ewert A, El Bihari S. 1971. Rapid recovery of Brugiamalayi following experimental infection of cats. Trans Royu Soc Trop Med Hyg 65;364-368.

Ewert A, Emerson GA. 1975. Effects of diethylcarbamazine on third stage Brugia malayi larvae in cats. Am J Trop Med Hyg 24:71-73.

Ewert A, Emerson GA. 1979. Effects of diethylcarbamazine citrate on fourth stage and adult Brugiamalayi in cats. Am J Trop Med Hyg 28:496-499.

Ewert A, Reitmeyer JC, Folse D. Chronic infection of cats with Brugiamalayi and streptococcus. SE Asian J Trop Med Publ Hlth 11:32-39.

Fader RC, Ewert A. 1986. Evolution of lymph thrombi in experimental Brugiamalayi infections: a scanning electron microscopic study. Lymphology 19:146-152.

Feng LC. 1936. The development of Microfilariamalayi in A. hyrcanus var. sinensis Wied. Chin Med J (Suppl) 1:345-367.

Folse DS, Ewert A. 1988. Edema resulting from experimental filariasis. Lymphology 21:244-247.

Folse D, Ewert A, Reitmeyer JC. 1981. Light and electron microscopic studies of lymph vessels from cats infected with Brugiamalayi. SE Asian J Trop Med Publ Hlth 12:174-184.

Hillman GR, Westerfield L, Ewert A, Wang YX. 1983. Serum levels of a filaricide, diethylcarbamazine citrate, in cats following different routes of administration. SE Asian J Trop Med Publ Hlth 14:171-175.

Laing ABG. 1961. Influence of the animal host on the microfilarial periodicity of Brugiamalayi. Trans Roy Soc Trop Med Hyg 55:558.

Mak JW, Lam PLW, Noor Rain A, Suresh K. 1987. Chemoprophylactic studies with ivermectin against subperiodic Brugiamalayi infection in the leaf monkey, Presbytiscristata. J Helminthol 61:311-314.

Mak JW, Yen PKF, Lim KC, Ramiah N. 1980. Zoonotic implications of cats and dogs in filarial transmission in Peninsular Malaysia. Trop Geogr MEd 32:259-264.

Ottesen EA; Duke BO; Karam M; Behbehani K. 1997. Strategies and tools for the control/elimination of lymphatic filariasis. Bull World Health Organ 75: 491 503.

Palmieri JR, Masbar S, Purnomo Marwoto HA, Tirtokusumo S, Darwis F. 1985. The domestic cat as a host for Brugian filariasis in South Kalimantan (Borneo), Indonesia. J Helmintol 59:277-281.

Paniker KN, Arunachalam N, Kumar MNP, Prathibha J, Sabesan S. 1997. Efficacy of diethylcarbamazine-medicated salt for microfilaraemia of Brugiamalayi. Natl Med J India 10:275-276.

Partono F, Oemijati S, Hudojo, Joesoef A, Clarke MD, Durfee PT, Irving GS, Taylor J, Cross JH. 1977. Brugiamalayi in seven villages in sourth Kalimantan, Indonesia. SE Asian J Ttrop Med Publ Hlth 8:400-407.

Phantana S, Shutidamrong C, Chusattayanend W. 1987. Brugiamalayi in a cat from southern Thailand. Trans Roy Soc Trop Med Hyg 81:173-174.

Prasomsitti P, Mak JW, Sucharit P, Liew LM. 1983. Detection of antibodies in cats infected with filarial parasites by the indirect immunofluorescence technique. SE Asian J Trop Med and Pub Hlth. 14:353-356.

Rao SS, Maplestone PA. 1940. The adult of Microfilariamalayi Brug, 1927. Ind Med Gazette 75:159-160.

Sakamoto M. 1980. Changes in the lymphatic system of cats experimentally infected with Brugia. Trop Med 22:223-236.

Sivanandam S, Fredericks HJ. 1966. The “Innenkörper” in differentiation between the microfilariae of Brugia pahangi and B. malayi (sub-periodic form). Med J Malaya 20:3387-338.

Sucharis S, Sitthai W, Surathin K. 1992. Natural infection of Brugiamalayi in a child outside endemic area, Thailand. Mosq Borne Dis Bull 9:128-130.

Wharton RH, JFB Edeson, ABG Laing. 1958. Laboratory transmission of Wuchereriamalayi by mosquito bites. Trans Roy Soc Trop Med Hyg 52:288


Figure 4-50. Brugia malayi. Microfilaria from human, Hematoxylin stain.

Figure 4-51. Brugia malayi. Microfilaria from human, Giemsa stain.

Comments are closed.