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Clinical Microbiology Reviews, October 2007, p. 579-592, Vol. 20, No. 4
0893-8512/07/$08.00+0     doi:10.1128/CMR.00027-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Plasmodium malariae: Parasite and Disease

William E. Collins* and Geoffrey M. Jeffery

Centers for Disease Control and Prevention, National Center for Zoonotic, Vector Borne and Enteric Diseases, Division of Parasitic Diseases, Chamblee, Georgia 30341, and U.S. Public Health Service, Atlanta, Georgia

SUMMARY
INTRODUCTION
LIFE HISTORY
    Human Host
        Prepatent period.
        Fever.
        Parasitemia.
        Recrudescence.
    Preerythrocytic Stages
    Mosquito Host
DISTRIBUTION
LABORATORY DIAGNOSIS
    Preservation
SEROLOGIC STUDIES
MOLECULAR STUDIES
INFECTIONS IN CHIMPANZEES AND MONKEYS
PATHOLOGY
ULTRASTRUCTURE
RELATIONSHIP TO OTHER SPECIES
REFERENCES

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SUMMARY
 
A review of the life history of Plasmodium malariae, the quartan malaria parasite of humans, is presented. Much of the information is based on data obtained from induced infections in humans who were given malaria therapy for the treatment of neurosyphilis between 1940 and 1963. Prepatent periods (i.e., the time until the first day of parasite detection) fever episodes, and maximum parasitemias as a result of infection with P. malariae were obtained and are presented. Experimental and known vectors of the parasite are also discussed. Splenectomized chimpanzees and New World monkeys are readily infected and serve as sources of parasites and antigens for diagnostic and molecular studies. South American monkeys are naturally infected with a parasite known as Plasmodium brasilianum. This parasite appears to be P. malariae that has adapted from humans to grow in monkeys, probably within the last 500 years. Infection with P. malariae is associated with the production of immune complexes in the kidneys and the associated nephrotic syndrome. The essential lesions are a thickening of the glomerular basement membrane and endocapillary cell proliferation. Studies of monkeys infected with P. malariae indicate the same pathology as that demonstrated in humans.


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INTRODUCTION
 
Plasmodium malariae is a malaria parasite that causes a disease that has been recognized since the Greek and Roman civilizations over 2,000 years ago. Quartan, tertian, and semitertian patterns of fever in patients were described by the early Greeks. After the discovery by Alphonse Laveran in 1880 (75) that the causative agent for malaria was a parasite, detailed studies on these organisms commenced. The early detailed work of Golgi in 1886 demonstrated that in some patients there was a relationship between the 72-hour life cycle of development of the parasites and a similar periodicity of the paroxysm (chill and fever pattern in the patient), whereas in other patients there were 48-hour cycles of development (54). He came to the conclusion that there must be more than one species of malaria parasite responsible for these different patterns of cyclical infection.

Eventually, the different parasites were separated and given the names that they carry at the present time. In 1890, Grassi and Feletti (58) reviewed the available information and named P. malariae and P. vivax with the following statement: "C'est pour cela que nous distinguons, dans le genre Haemamoeba, trois espèces (H. malariae de la fièvre quarte, H. vivax de la fièvre tierce et H. praecox de la fièvre quotidienne avec coutres intermittences etc.)." The current name for the parasite that we discuss here is Plasmodium malariae (Grassi and Feletti 1890).


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LIFE HISTORY
 
Plasmodium malariae has developmental cycles in the mosquito and in the primate host (20). When gametocytes are ingested during mosquito feeding, a process called exflagellation of the microgametocyte occurs, resulting in the formation of up to eight mobile microgametes. Following fertilization of the macrogamete, a mobile ookinete is formed, which penetrates the peritropic membrane surrounding the blood meal and travels to the outer wall of the midgut of the Anopheles mosquito. There, under the basal membrane, the oocyst develops. After a period of 2 to 3 weeks, depending on the temperature, many hundreds to a few thousand sporozoites are produced within each oocyst. The oocyst ruptures and the sporozoites are released into the hemocoel of the mosquito. The sporozoites are carried by the circulation of the hemolymph to the salivary glands, where they become concentrated in the acinal cells. During feeding, a small number of sporozoites (<100) are introduced into the salivary duct and injected into the venules of the bitten human, to initiate the cycle in the liver.

In the human, following introduction into the bloodstream, the sporozoites rapidly invade the liver within an hour, where, within a parenchymal cell, the parasite matures in approximately 15 days. Eventually many thousands of merozoites are produced in each schizont. Upon release, these merozoites invade erythrocytes and initiate the erythrocytic cycle. There is no evidence of quiescent liver stage forms (hypnozoites) such as are found in P. vivax and P. ovale infections in humans. However, not all liver stage forms will mature on the same day; biopsies indicate that these forms may rupture and release parasites over a number of days. Following a developmental cycle in the erythrocyte that lasts, on average, for 72 h, from 6 to 14 (average, 8) merozoites are released to reinvade other erythrocytes. Some of the merozoites develop into the two forms of gametocytes (micro- and macrogametocytes). When they are taken into the mosquito during feeding, the cycle is repeated.

Human Host

Prepatent period. There are only a limited number of reports on the transmission of P. malariae to humans to determine prepatent periods. The prepatent period is defined as the time until the first day that parasites are detected on a thick blood film. Shute and Maryon (104) reported the shortest prepatent period of 16 days for a West African strain. Boyd and Stratman-Thomas (10) reported prepatent periods of 27, 32, and 37 days for two different strains, and Mer (86) transmitted a Palestinian strain to three patients, in whom the periods were 26, 28, and 31 days. Prepatent periods of 23 and 26 days were reported by de Buck (42) for four patients infected with a Vienna strain, and Boyd and Stratman-Thomas (12) reported 28- and 40-day prepatent periods. Marotta and Sandicchi (81) reported incubation periods (days until symptoms first appeared) of 23 and 29 days in two patients. Boyd (9) reported on three different strains for which prepatent periods ranged from 28 to 37 days. Siddons (107) reported a prepatent period of 30 days, and Young and Burgess (120) reported prepatent periods of 29 and 59 days. Mackerras and Ercole (79) reported a 24-day period for a Melanesian strain, and Kitchen (72) reported a mean prepatent period of 32.2 days (range, 27 to 37 days) for American strains of P. malariae. Young and Burgess (121) transmitted the USPHS strain of P. malariae to patients, and the prepatent periods were 33 and 36 days. Ciuca et al. (18) reported prepatent periods for the Romanian VS strain ranging from 18 to 25 days. Lupascu et al. (78) reported incubation periods of 18 to 19 days for the VS strain; in four additional patients the prepatent period ranged from 21 to 30 days (48). In transmission studies with a Nigerian strain involving four volunteers, the prepatent periods ranged from 24 to 33 days (40). Thus, as these data show, there is a wide range in the length in the prepatent period in mosquito-transmitted P. malariae (16 to 59 days).

Fever. The most detailed study of the paroxysm of P. malariae is probably that by Young et al. (123) in which they examined 420 paroxysms. The average fever peak was 104.1°F (rectally), with the highest recorded being 106.4°F. Fevers (≥101°F) ranged in duration from 5 to 32 h, with an average of 10 h 58 min. Some fevers were introduced by chills, while others were not.

A retrospective examination of induced infections with P. malariae was made by McKenzie et al. (83). These data were extracted from the records of patients who were given malaria therapy for the treatment of neurosyphilis between 1940 and 1963. Prior to the introduction of penicillin for the treatment of syphilis, malaria was one of the most effective treatments for the disease (118). It was estimated that perhaps 20% of patients in U.S. mental hospitals had neurosyphilis (62), and infection with P. vivax or P. malariae was standard practice in the treatment of the disease. Plasmodium falciparum was less commonly used because of the difficulty of controlling infections with this species of parasite. It was believed that a combination of repeated episodes of high-intensity fever combined with a nonspecific stimulation of the immune system induced by the malaria parasite combined to destroy the spirochete. Because most African American patients were resistant to infection with P. vivax (due to the Duffy negative blood grouping), they were most often treated with P. malariae.

A listing of the days of fever of ≥101°F and ≥104°F and the maximum fevers for 69 of the patients examined by McKenzie et al. (83) with no known previous malaria infection who were allowed to have parasitemia of P. malariae for 60 or more days is presented in Table 1. For these patients, the median number of days of fever of ≥101°F was 21.9 and the median number of days of fever of ≥104°F was 10.2. The median maximum fever for the 69 patients was 105.6°F. One patient (S-1112) failed to exhibit fever of ≥101°F in spite of a maximum parasite count of 4,100/µl. Fever often occurred on an every-third-day pattern, as shown in Fig. 1. It is also apparent that the fever occurred just prior to an increased parasite count associated with release of a new population of parasites. Because fever regularly occurs again on the fourth day in many patients, P. malariae infections are often referred to as being "quartan" malaria.


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  		TABLE 1. Parasite counts and fever for 69 patients infected with Plasmodium malariae for 60 days or morea


Figure 1
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  		FIG. 1. Daily peak parasite counts and fever peaks in a patient infected with Plasmodium malariae, showing the synchronous quartan pattern of fever and peak parasite count.

Parasitemia. Maximum parasite counts are usually low compared to those in patients infected with P. falciparum or P. vivax. This is due to several factors: (i) the lower number of merozoites produced per erythrocytic cycle, (ii) the extended 72-hour developmental cycle versus the 48-hour cycle of P. vivax and P. falciparum, (iii) the preference of the parasite to developed in older erythrocytes, and (iv) the combination of these factors that allows for the earlier development of immunity by the human host. In the 69 patients (Table 1), the maximum parasite count ranged from 1,648/µl to 49,680/µl, with a median count of 8,875/µl (10,000/µl = 0.25% of erythrocytes infected). Some patients had long periods of parasitemia and extended periods when parasite counts were >1,000/µl. These patients averaged 50.5 days with parasite counts of >1,000/µl. When the parasite counts for these patients were averaged for the first 40 days of patent parasitemia (Fig. 2), it was apparent that the parasite count peaked at approximately 2 weeks and then remained relatively stable. The median parasite count actually did not begin to decline until 60 days or more of patent parasitemia.


Figure 2
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  		FIG. 2. Median parasite counts during the first 40 days of patent parasitemia for 69 patients infected with Plasmodium malariae. Maximum parasite counts are limited in infections with P. malariae due to the low number of merozoites produced, 72-hour developmental cycle, and preference for older erythrocytes.

Other patients in the data studied by McKenzie et al. (83) had been infected with P. malariae following previous infection with other species of malaria parasites. Forty-six patients were infected following infection with P. falciparum (Table 2). The maximum parasite counts ranged from 312/µl to 29,825/µl, with the median being 6,608/µl. The length of parasitemia was shorter, and there were fewer days with parasite counts of >1,000/µl. The ratio between the number of days of fever of ≥101°F to ≥104°F was almost identical to that for patients with no previous infection. In addition, 39 patients had been infected with P. malariae following infection with P. vivax (Table 3). The maximum parasite counts ranged from 424/µl to 19,624/µl, with a median of 9,250/µl. Only eight patients were infected with P. malariae following infection with P. ovale (Table 4). The median maximum parasite count was 13,575/µl.


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  		TABLE 2. Parasite counts and fever for 46 patients infected with Plasmodium malariae following previous infection with P. falciparuma


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  		TABLE 3. Parasite counts and fever for 39 patients infected with Plasmodium malariae following previous infection with P. vivaxa


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  		TABLE 4. Parasite counts and fever for eight patients infected with Plasmodium malariae following previous infection with P. ovalea

Recrudescence. Plasmodium malariae does not relapse from persistent liver stage parasites. However, the blood stage parasites persist for extremely long periods, often, it is believed, for the life of the human host. There have been many reports of people who have left zones of endemicity and, either following donation of blood in which the recipient developed an infection or under stress, whose infections have recrudesced after many years of dormancy. For example, Collins et al. (33) reported on a transfusion case in which the donor had probably acquired infection with P. malariae in China 50 years previously. Vinetz et al. (116) report a case of an infection acquired in Greece over 40 years (and possibly up to 70 years) previous to splenectomy and subsequent diagnosis. Because almost all of these long-term infections have been detected following transfusion donations, it is believed that the parasites have persisted in the blood at very low densities.

Preerythrocytic Stages

The preerythrocytic tissue stages develop in the liver following the introduction of sporozoites. The time required for maturation and release of merozoites from the mature schizonts to invasion of erythrocytes is approximately 15 days. The tissue stages of P. malariae were first described by Bray (13, 14) in liver biopsy specimens from sporozoite-inoculated chimpanzees. The host cell nucleus was enlarged and pushed to one side. In over 50% of the parasitized parenchymal cells, two or more nuclei were present. He was able to describe the 8-, 9-, 10-, 11-, 12-, and 12.5-day-old forms. The nuclei were always randomly distributed; there were no pseudocytomeres, no evidence of septum formation or plasmotomy, and no mature schizonts at these time points.

Lupascu et al. (77) obtained biopsy material from a chimpanzee at 12, 13, 14, and 15 days after introduction of sporozoites of P. malariae. The schizonts were considered mature at 15 days. The main characteristics were enlargement of the host cell nucleus, many peripheral and internal vacuoles, no cytomeres, large clefts, red-staining strands, and plaques in the mature schizonts.

Millet et al. (87) reported the development of preerythrocytic stages of P. malariae in cultures of hepatocytes from chimpanzees and Aotus lemurinus griseimembra monkeys. Schizonts were observed in chimpanzee hepatocytes at 8, 11, and 13 days after inoculation of sporozoites. Only one schizont was seen in Aotus hepatocytes at day 13.

Mosquito Host

Many different vectors have been shown to be capable, at least experimentally, of infection with this parasite. These are listed in Table 5. Those that have been proven to be capable of transmitting P. malariae to humans experimentally are also indicated. The development of P. malariae in mosquitoes has been described by a number of workers; the first definitive studies were carried out by Shute and Maryon (105) on its development in Anopheles atroparvus mosquitoes. In the studies of Collins et al. (38) with Anopheles freeborni, when incubated at a temperature of 25°C, sporozoites were present in the salivary glands in 17 days. At day 6, the mean oocyst diameter was 12 µm, with a range of 9 to 14 µm. The oocysts continued to grow so that by day 14 they ranged from 20 to 65 µm, with a mean of 38 µm. Early differentiation and formation of sporozoites were apparent by day 14 (Fig. 3).


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  		TABLE 5. Species of Anopheles mosquitoes that have been infected with Plasmodium malariae


Figure 3
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  		FIG. 3. Development of oocysts of Plasmodium malariae in Anopheles freeborni mosquitoes. Top row, 10-, 11-, 12-, and 13-day oocysts; bottom row, 14-, 15-, and 17-day oocysts and sporozoites.


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DISTRIBUTION
 
In general, the distribution of P. malariae coincides with that of P. falciparum. In areas of endemicity in Africa, infections of P. malariae are mixed with P. falciparum infections. In many instances, the presence of P. malariae infections is unapparent unless PCR techniques are used to reveal low-level or subpatent infections. Plasmodium malariae is wide spread throughout sub-Saharan Africa, much of southeast Asia, into Indonesia, and on many of the islands of the western Pacific. It is also reported in areas of the Amazon Basin of South America, along with Plasmodium brasilianum, a parasite commonly found in New World monkeys. This parasite is apparently the same species as P. malariae that has naturally adapted to grow in monkeys following human settlement of South America within the last 500 years. In the recent past, P. malariae was prevalent in Europe and in southern parts of the United States.


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LABORATORY DIAGNOSIS
 
Diagnosis of P. malariae infection is preferentially made by the examination of peripheral blood films stained with Giemsa stain. PCR techniques are now routinely used in many laboratories to confirm diagnoses and to separate mixed infections. Recently, in southeast Asia it has been shown that infections with the monkey malaria parasite Plasmodium knowlesi in humans have been misdiagnosed as being infections with P. malariae (68, 108). Identification was confirmed by PCR. Thus, careful microscopic examination may not be sufficient for positive confirmation in certain situations where monkey malaria parasites such as P. knowlesi or P. inui may be transmitted to humans. In areas of South America where humans and monkeys coexist, it is impossible to differentiate infections of P. malariae from infections of P. brasilianum because they may, in fact, be one and the same.

The first stages that appear in the blood are the ring stages that are formed by the invasion of merozoites released by rupturing liver stage schizonts. As described by Coatney et al. (Fig. 4) (20), these grow slowly but soon occupy one-fourth to one-third of the parasitized cell. Pigment increases rapidly, and the half-grown parasite may have from 30 to 50 jet-black granules. As the parasite grows, it assumes various shapes, and it often stretches across the host cell to form what is known as the band form. These are often considered diagnostic, although they are sometimes seen in other species. The host cell is not enlarged as the parasite grows to fill the infected erythrocyte.


Figure 4
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  		FIG. 4. Development of the erythrocytic stages of Plasmodium malariae. 1, normal red cell; 2 to 5, young trophozoites; 6 to 11, growing trophozoites; 12 and 13, nearly mature and mature trophozoites, respectively; 14 to 20, developing schizonts; 21 and 22, mature schizonts; 23, developing gametocyte; 24, mature macrogametocyte; 25, mature microgametocyte. (Reprinted from reference 20.)

At about the 54th hour, segmentation begins, and by the 65th hour, the host cell is nearly filled and the parasite contains five or six chromatin masses; pigment is scattered. The nuclei and cytoplasm begin to separate, and the pigment becomes segregated and clumped in a loose mass in the center of the cell surrounded by the more or less symmetrically arranged merozoites. The number of merozoites may be from 6 to 14, but the average number is 8.

The mature macrogametocyte has a dense, deeply staining blue cytoplasm with a small red-staining nucleus. The pigment is scattered. The parasite completely fills the host cell. The cytoplasm of the adult microgametocyte has a light bluish pink stain. The pigment is limited to the cytoplasm of the parasite. The nucleus is diffuse, takes a pinkish-blue stain, and may occupy half the infected cell. The parasite appears to occupy the entire host cell. Ordinarily, microgametocytes outnumber the macrogametocytes.

Snounou et al. (109) applied the nested PCR technique to the diagnostic identification of all four human-infecting species of Plasmodium, using genus- and species-specific primers targeting the 18S rRNA gene. Failure to detect some P. malariae infections has prompted alteration of the species-specific primers for this parasite.

Recent efforts have been directed towards the development of real-time PCR assays. Rougemonet et al. (97) used a set of generic primers targeting a highly conserved region of the 18S rRNA genes of the four human-infecting species of Plasmodium to develop such an assay, which was highly specific and sensitive.

McNamara et al. (85) described a PCR/ligase detection reaction fluorescent-microsphere assay for the diagnosis of infection levels with all four species of human malaria, which shows promise for the detection of minority species in infections with mixed Plasmodium species.

Preservation

The preservation of viable malaria parasites by freezing made possible the study of these organisms without continuous cyclical passage. In 1955, Jeffery and Rendtorff (67) reported the frozen preservation of blood stages of P. malariae. Blood stages were stored for 20 and 60 days at a temperature of –70°C. The frozen preservation of P. malariae-infected erythrocytes has now become routine. Once the infections were established in chimpanzees and New World monkeys, subsequent infections were most frequently induced by the injection of parasitized erythrocytes that had been stored frozen over liquid nitrogen, often after many years of storage. Parasites are usually stored in Glycerolyte (Baxter Healthcare Corp., Fenwal Div., Deerfield, IL) and are expected to be viable for decades when held at extremely low temperatures over liquid nitrogen. Thick and thin blood films for immunofluorescence studies and teaching can be stored unfixed and frozen for extended periods. However, frozen blood is unsuitable for the preparation of blood films for microscopic diagnosis.


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SEROLOGIC STUDIES
 
Serologic tests are not specific enough for diagnostic purposes but are basic epidemiologic tools. They allow for the measurement of past exposure to infection. The immunofluorescent-antibody (IFA) technique has been used to measure the presence of antibodies to P. malariae. It was shown that when an infection was of short duration, the response soon declined. However, if the parasite count recrudesced or reinfection occurred, the IFA response rose to a higher lever and persisted for many months or years, as shown in Fig. 5 (27). Cross-reaction studies indicated that P. brasilianum, the monkey malaria parasite from South American monkeys that appears to be identical to P. malariae, could be used in serologic testing (26). Plasmodium fieldi, a parasite of macaques from southeast Asia, also cross-reacted strongly with P. malariae (26). In a serologic study of 498 sera collected from Nigerians, 43.2% had positive responses to P. brasilianum (35). The response was low in children but was equal to that to P. falciparum with sera from individuals 13 years of age and older. In a study of a jungle aboriginal area in Malaysia, there was an almost complete absence of P. malariae infection during a parasitologic survey, whereas historically the incidence was known to be quite high (38). The high incidence of maximum IFA responses (51%) to P. malariae, however, was probably more indicative of the malarial experience or of subpatent parasitemia than the slide survey because of recent drug interventions (38).


Figure 5
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  		FIG. 5. Development of the IFA response in a patient following infection and recrudescence of an infection with Plasmodium malariae. Recrudescence of infection began at day 84.

The structure of the circumsporozoite (CS) gene of P. malariae was first described by Lal et al. (74). Serologic studies were subsequently conducted for responses to CS proteins of P. malariae by using the CS repeat (NAAG)5 in an enzyme-linked immunosorbent assay (ELISA). In a study in Asembo Bay, Kenya, 59% of persons had antibodies to the peptide; all positivity rates increased with age (43). In a seroepidemiologic study conducted on Indian tribes in the Amazon Basin of northern Brazil, Arruda et al. (41) found that almost all Metuktire and almost 90% of the Asurini adults had antisporozoite antibodies against P. brasilianum/P. malariae. A monoclonal antibody specific for the repeat epitope of the CS protein of P. malariae was developed to detect sporozoites in infected mosquitoes (22). The (NAAG)5 ELISA has also been used extensively. Beier et al. (7), for example, identified 3.2% of infected Anopheles gambiae sensu lato and A. funestus mosquitoes collected in western Kenya as being infected with P. malariae. This has proven to be a valuable epidemiologic tool in identifying potential vectors of P. malariae.


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MOLECULAR STUDIES
 
Cochrane et al. (21) produced a hybridoma secreting a monoclonal antibody against the CS protein of P. malariae (Uganda I/CDC strain). A two-dimensional electrophoretic assay showed that the CS protein recognized by the monoclonal antibody contains a repetitive epitope. The antibody also reacted strongly with sporozoites of the simian parasite P. brasilianum but did not bind to sporozoites of P. falciparum, P. vivax, and P. ovale. Monoclonal antibodies specific for a repeat epitope of the CS protein of P. malariae sporozoites were then used to develop a two-site, single-antibody-based ELISA to detect sporozoites in mosquitoes (22). The major repeat was determined to be Asn Ala Ala Gly (NAAG), with two different minor repeats, Asn Asp Ala Gly (NDAG) and Asn Asp Gln Gly (NDEG). In a study in Cameroon, the length of the CS protein gene varied due to the number of tandem repeat units (115).

A gene encoding the small-subunit rRNA of P. malariae was sequenced and shown to contain unique regions that could be used as diagnostic probes (55). Studies indicated a variant form in the small-subunit rRNA gene sequence in the Sichuan province of China and along the Thai-Myanmar border, by deletion of 19 bp and seven substitutions of base pairs in the target sequence (76). Thus, there appear to be two different types or potentially two subspecies of P. malariae, based on molecular differences in Asian parasites.

There are few genomic data on this parasite. Studies on the gene encoding cytochrome b from the linear mitochondrial genome indicated that P. malariae was separate from other members of the primate-infecting Plasmodium species (46). Plasmodium inui and P. malariae do not form a monophyletic group, demonstrating that periodicity is convergent in the evolution of the genus.


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INFECTIONS IN CHIMPANZEES AND MONKEYS
 
Attempts to infect Old World monkeys have been unsuccessful. The first adaptation of P. malariae to New World monkeys was reported by Geiman and Siddiqui (49). Additional studies were made with different species of Aotus and Saimiri monkeys (23, 28, 32, 33, 34, 36). In splenectomized Aotus monkeys, maximum parasite counts with P. malariae varied markedly, from 10 to 56,800/µl. Parasitemia often persisted for many weeks; recrudescence occurred, and mosquito infection was readily obtained (Fig. 6). The median maximum parasite count depended on the previous heterologous malarial experience of the animals. When 18 Aotus monkeys with no previous history of infection were infected with P. malariae, the median maximum parasite count was 13,760/µl. In 29 monkeys that had been previously infected with P. falciparum, the median maximum parasite count was 6,270/µl. In 46 monkeys that had been previously infected with P. vivax, the maximum parasite count with P. malariae was 1,488/µl. Following the infection of 49 animals that had been previously infected with both P. vivax and P. falciparum, the median maximum parasite count with P. malariae was only 899/µl. Splenectomized Saimiri boliviensis monkeys had maximum parasite counts that varied from 62/µl to 22,134/µl.


Figure 6
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  		FIG. 6. Daily parasite counts and percent infection of Anopheles freeborni mosquitoes when fed on a splenectomized Aotus lemurinus griseimembra monkey infected with the Uganda I strain of Plasmodium malariae.

Splenectomized chimpanzees were shown by Rodhain (95) and Garnham et al. (48) to be readily infected. Bray (14) observed parasite counts in splenectomized animals of between 25,000 and 50,000 per µl, and Garnham et al. (48) observed a maximum parasite count of 160,000 per µl. In a study with 31 splenectomized chimpanzees with various previous histories of infection with P. vivax and P. ovale, maximum parasite counts following inoculation with a strain of P. malariae from Uganda ranged from 930 to 75,700 per µl (29). Infections were infective to a variety of mosquito species on more than 50% of the days on which they were fed. In most instances, infection was obtained when the parasite count was rising and diminished as soon as the count peaked.

In 1920, Reichenow (91) studied malarias in chimpanzees and gorillas in the Cameroons and found P. malariae. Blacklock and Adler (8) in 1922 in Sierra Leone and Schwetz (100, 101, 102) in the Belgian Congo also saw P. malariae in these animals. In 1939, Brumpt (15) gave the name P. rodhaini to the quartan parasite that infected chimpanzees and gorillas. However, subsequent transmission studies with quartan parasites isolated from chimpanzees convinced investigators that this parasite was actually P. malariae (92, 93, 94, 96).


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PATHOLOGY
 
Watson in 1905 (117) noted the presence of edema in a patient with malaria in Malaysia, and subsequently the relationship between P. malariae infection and the nephrotic syndrome has been well documented. Many investigators (9, 51, 52, 53, 110) indicated a close relationship between quartan malaria and renal disease. Hendrikse and Adeniyi (60) described the clinico-pathological features associated with infection with P. malariae and suggested that immune complexes may cause structural glomerular damage. Dixon (44) demonstrated immune complexes in the kidneys of patients with nephrotic syndrome associated with quartan malaria.

The essential lesions are a thickening of the glomerular basement membrane and endocapillary cell proliferation (61, 71). This gives rise to a double-contour or plexiform arrangement of periodic acid-Schiff stain-positive, argyrophilic fibrils (45, 61, 119). As the disease progresses, more capillaries become affected, and the lesions extend to cause progressive narrowing and eventually complete obliteration of capillary lumens.

Electron microscopy shows thickening of the glomerular basement membrane with an increase in the basement membrane-like material of varying density in the subendotherial zone (2). Hendrickse et al. (61) graded the severity of pathological changes based on the percentage of glomeruli showing lesions. If patients had up to 30% of glomeruli showing lesions, they responded to therapy. However, if they had greater than that, they did not show a response to therapy. The renal disease tended to become chronic and nonresponsive to treatment with antimalarial and immunosuppressive drugs.

Aikawa et al. (1) examined the kidneys of Aotus monkeys infected with P. malariae and demonstrated that the nephrotic syndrome seen in monkeys was consistent with that seen in humans. Histologically, glomeruli of these monkeys infected with P. malariae showed thickening and reduplication of the basement membrane and endocapillary cell proliferation. Electron microscopy revealed electron-dense deposits in the subendothelial and mesangial areas. The changes were consistent with membranoproliferative glomerulonephritis, similar to that of humans infected with P. malariae.


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ULTRASTRUCTURE
 
Ultrastructural studies have also been made on the erythrocytic stages of P. malariae and on the oocyst and sporozoite stages in the mosquito. Atkinson et al. (4) indicated that P. malariae was morphologically indistinguishable and structurally similar to other primate malaria species. There were highly structured arrays of merozoite surface coat proteins in the cytoplasm of early schizonts and on the surface of budding merozoites. Knobs were present in the membranes of Maurer's clefts. Morphological evidence suggested that proteins are transported between the erythrocyte surface and intracellular parasites via two routes: one associated with Maurer's clefts for transport of membrane-associated knob material and a second associated with caveolae in the host cell membrane for the import or export of host- or parasite-derived substances through the erythrocyte cytoplasm.

Nagasawa et al. (90) used immunoelectron microscopy and a monoclonal antibody for the CS protein of P. malariae to determine ultrastructural localization of this protein in midgut oocysts and salivary gland sporozoites. The CS protein was found along the capsule of immature oocysts but rarely within the cytoplasm. It was detected on the inner surface of peripheral vacuoles during oocyst maturation and on the plasma membrane of the sporoblast. Salivary gland sporozoites and sporozoites in mature oocysts were labeled uniformly on the outer surfaces of their plasma membranes. Antibodies against P. brasilianum CS protein reacted with P. malariae sporozoites.


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RELATIONSHIP TO OTHER SPECIES
 
Malaria parasites of primates are organized based on biologic characteristics. Plasmodium brasilianum, the monkey-infecting malaria parasite of South America, probably is an adaptation of P. malariae to New World primates with the introduction of Old World humans to the New World. The adaptation probably occurred within the last 500 years with the introduction of large numbers of people from Africa, where P. malariae was prevalent. The chronic nature of the parasite easily allowed for its survival in the human host during transit to the New World. The ready passage of P. brasilianum to humans and the passage of P. malariae to New World monkeys indicated that such interspecies transmission between primates and humans is both feasible and probable.

In southeast Asia, the other complex of parasites with a 72-hour developmental cycle in the blood of the primate host is Plasmodium inui. This parasite is also experimentally transmissible to humans (19). Whether or not such transmission occurs in nature has not been demonstrated. However, monkeys are commonly found to be infected with P. inui in close proximity to humans, and many different mosquito vectors are capable of transmitting the parasite to humans. Morphologically, it would be difficult to separate infections with monkey malaria parasites such as P. knowlesi and P. inui from those with P. malariae, particularly if reliance on a thick blood film diagnosis was made. This is illustrated in Fig. 7, which shows the blood stages of P. malariae, P. inui, and P. knowlesi. Neither of the erythrocytes that are infected with trophozoites of these parasites shows cellular enlargement or prominent stippling. Initial examination of the blood film, if from a human, would have immediately ruled out infection with P. vivax and P. ovale, both of which result in enlargement of the host cell erythrocyte and prominent stippling, or with P. falciparum, which rarely exhibits mature forms in the peripheral blood. Thus, the diagnostician would be left with P. malariae as the probable diagnosis. Only the proximity of monkeys would have suggested a secondary examination by PCR or subpassage to susceptible monkeys to confirm infection with a Plasmodium species other than P. malariae.


Figure 7
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  		FIG. 7. Top row, trophozoite stages of Plasmodium malariae; middle row, trophozoite stages of Plasmodium knowlesi; bottom row, trophozoite stages of Plasmodium inui.

There is no molecular evidence suggesting that P. malariae is closely related to any of the other primate malaria parasites that have been thus far examined (except P. brasilianum). Plasmodium malariae and P. brasilianum are either the same species or variants of the same species. Plasmodium malariae appears to represent an independent colonization of humans by malaria parasites (46). However, there is no indication of a close relationship to other primate-infecting species of Plasmodium, and the evolutionary origin of the species is unclear.


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FOOTNOTES
 
* Corresponding author. Mailing address: Centers for Disease Control and Prevention, National Center for Zoonotic, Vector Borne and Enteric Diseases, Division of Parasitic Diseases, Chamblee, GA 30341. Phone: (770) 488-4077. Fax: (770) 488-4253. E-mail: wec1{at}cdc.gov Back


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REFERENCES
 
    1
  1. Aikawa, M., J. R. Broderson, I. Igarashi, G. Jacobs, M. Pappaioanou, W. E. Collins, and C. C. Campbell. 1988. An atlas of renal disease in Aotus monkeys with experimental plasmodial infection, p. 1-97. American Institute of Biological Sciences, Washington, DC.
    2. 2
  2. Allison, A. C., V. Houba, R. G. Hendrickse, S. De Petrise, G. M. Edington, and A. Adeniyi. 1969. Immune complexes in the nephritic syndrome of African children. Lancet 1:1232-1238.[Medline]
    3. 3
  3. Anazawa, K. 1931. Experimental studies on the infectibility of anopheline mosquitoes of Formosa. J. Med. Assoc. Formosa 30:609-632.
    4. 4
  4. Atkinson, C. T., M. Aikawa, A. P. Rock, K. Marsh, P. M. Andrysiak, G. H. Campbell, W. E. Collins, and R. J. Howard. 1987. Ultrastructure of the erythrocytic stages of Plasmodium malariae. J. Protozool. 34:267-274.[Medline]
    5. 5
  5. Averbouch, I. 1930. Expériences sur la transmission á l'homme de la fiévre quarte par les piqûres d'Anopheles maculipennis var saccharovi. Rev. Microbiol. Epid. Parasit. 9:379-381.
    6. 6
  6. Basu, B. C. 1943. Laboratory studies on the infectivity of Anopheles annularis. J. Malaria Inst. India 5:31-51.
    7. 7
  7. Beier, M. S., I. K. Schwartz, J. C. Beier, P. V. Perkins, F. Onyango, J. K. Koros, G. H. Campbell, P. M. Andrysiak, and A. D. Brandling-Bennett. 1988. Identification of malaria species by ELISA in sporozoite and oocyst infected Anopheles from western Kenya. Am. J. Trop. Med. Hyg. 39:323-327.[Abstract/Free Full Text]
    8. 8
  8. Blacklock, B., and S. Adler. 1922. A parasite resembling Plasmodium falciparumin a chimpanzee. Ann. Trop. Med. Parasitol. 16:99-106.
    9. 9
  9. Boyd, M. F. 1940. Observations on naturally and artificially induced quartan malaria. Am. J. Trop. Med. 20:749-798.
    10. 10
  10. Boyd, M. F., and W. K. Stratman-Thomas. 1933. A note on the transmission of quartan malaria by Anopheles quadrimaculatus. Am. J. Trop. Med. 13:265-271.
    11. 11
  11. Boyd, M. F., and W. K. Stratman-Thomas. 1934. The comparative susceptibility of Anopheles quadrimaculatus Say, and of Anopheles crucians Wied. (inland variety) to the parasites of human malaria. Am. J. Hyg. 20:247-257.
    12. 12
  12. Boyd, M. F., and W. K. Stratman-Thomas. 1936. The transmission of quartan malaria through two consecutive human-anopheline passages. Am. J. Trop. Med. 16:63-65.
    13. 13
  13. Bray, R. S. 1959. Pre-erythrocytic stages of human malaria parasites: Plasmodium malariae. Br. Med. J. 2:679-680.[Free Full Text]
    14. 14
  14. Bray, R. S. 1960. Studies on malaria in chimpanzees. VIII. The experimental transmission and pre-erythrocytic phase of Plasmodium malariae, with a note on the host-range of the parasite. Am. J. Trop. Med. Hyg. 9:455-465.[Abstract/Free Full Text]
    15. 15
  15. Brumpt, E. 1939. Les parasites du paludisme des chimpanzés. C. R. Soc. Biol. 130:837-840.
    16. 16
  16. Brumpt, E. 1949. The human parasites of the genus Plasmodium, p. 65-121. In M. F. Boyd (ed.), Malariology, vol. I. W. B. Saunders Co., Philadelphia, PA.
    17. 17
  17. Cambournac, F. J. C. 1942. Sôbre a epidemiologia do sezonismo em Portugal. Sociedade Industrial de Tipografia, Lda., Lisbon, Portugal.
    18. 18
  18. Ciuca, M., G. Lupascu, E. Negulici, and P. Constantinescu. 1964. Recherches sur la transmission expérimentale de P. malariae à l'homme. Arch. Roum. Pathol. Exp. Microbiol. 23:763-776.[Medline]
    19. 19
  19. Coatney, G. R., W. Chin, P. G. Contacos, and H. K. King. 1966. Plasmodium inui, a quartan-type malaria parasite of Old World monkeys transmissible to man. J. Parasitol. 52:660-663.[CrossRef][Medline]
    20. 20
  20. Coatney, G. R., W. E. Collins, M. Warren, and P. G. Contacos. 1971. The primate malarias. National Institutes of Health, Bethesda, MD.
    21. 21
  21. Cochrane, A. D., W. E. Collins, and R. S. Nussenzweig. 1984. Monoclonal antibodies identify the circumsporozoite protein of Plasmodium malariae and detect a common epitope on P. brasilianum sporozoites. Infect. Immun. 45:592-595.[Abstract/Free Full Text]
    22. 22
  22. Collins, F. H., P. M. Procell, G. H. Campbell, and W. E. Collins. 1988. Monoclonal antibody-based enzyme-linked immunosorbent assay (ELISA) for detection of Plasmodium malariae sporozoites in mosquitoes. Am. J. Trop. Med. Hyg. 38:283-288.[Abstract/Free Full Text]
    23. 23
  23. Collins, W. E., and P. G. Contacos. 1969. Infectivity of Plasmodium malariae in the Aotus trivirgatus monkey to Anopheles freeborni mosquitoes. J. Parasitol. 55:1253-1257.[CrossRef][Medline]
    24. 24
  24. Collins, W. E., P. G. Contacos, and W. Chin. 1973. Experimental infection in man with Plasmodium malariae. Am. J. Trop. Med. Hyg. 22:685-692.[Medline]
    25. 25
  25. Collins, W. E., P. G. Contacos, and B. B. Richardson. 1975. Studies on human malaria in Aotus monkeys. VI. Infectivity of Plasmodium malariae to different anophelines. J. Parasitol. 61:941-943.[CrossRef][Medline]
    26. 26
  26. Collins, W. E., G. M. Jeffery, E. Guinn, and J. C. Skinner. 1966. Fluorescent antibody studies in human malaria. IV. Cross-reactions between human and simian malaria. Am. J. Trop. Med. Hyg. 15:11-15.[Abstract/Free Full Text]
    27. 27
  27. Collins, W. E., G. M. Jeffery, and J. C. Skinner. 1964. Fluorescent antibody studies in human malaria. I. Development of antibodies to Plasmodium malariae. Am. J. Trop. Med. Hyg. 13:l-5.
    28. 28
  28. Collins, W. E., H. O. Lobel, H. McClure, E. Strobert, G. G. Galland, F. Taylor, A. L. Baretto. R. R. Roberto, J. C. Skinner, S. Adams, C. L. Morris, and J. Sullivan. 1994. The China I/CDC strain of Plasmodium malariae in Aotus monkeys and chimpanzees. Am. J. Trop. Med. Hyg. 50:28-32.[Abstract/Free Full Text]
    29. 29
  29. Collins, W. E., H. M. McClure, E. Strobert, V. K. Flipski, J. C. Skinner, P. S. Stanfill, B. B. Richardson, and C. Morris. 1990. Infection of chimpanzees with the Uganda I/CDC strain of Plasmodium malariae. Am. J. Trop. Med. Hyg. 42:99-103.[Abstract/Free Full Text]
    30. 30
  30. Collins, W. E., H. McClure, E. Strobert, B. B. Richardson, B. B. Richardson, J. M. Roberts, G. G. Galland, J. Sullivan, C. L. Morris, and S. R. Adams. 1993. Experimental infection of Anophele gambiae s. s. with Plasmodium malariae and P. brasilianum. J. Am. Mosq. Control Assoc. 9:68-71.[Medline]
    31. 31
  31. Collins, W. E., B. B. Richardson, J. S. Sullivan, C. L. Morris, and G. G. Galland. 1997. Infection of Anopheles freeborni mosquitoes on New World monkeys infected with the Uganda I/CDC strain of Plasmodium malariae. J. Parasitol. 83:1099-1103.[CrossRef][Medline]
    32. 32
  32. Collins, W. E., I. K. Schwartz, J. C. Skinner, and J. R. Broderson. 1984. Studies on the Uganda I/CDC strain of Plasmodium malariae in Bolivian Aotus monkeys and different anophelines. J. Parasitol. 70:677-681.[CrossRef][Medline]
    33. 33
  33. Collins, W. E., J. C. Skinner, J. R. Broderson, M. Pappaioanou, V. Filipski, B. B. Sutton, P. S. Stanfill, A. Y. Huong, J. Roberts, and C. Wilson. 1989. The Uganda I/CDC strain of Plasmodium malariae in Aotus lemurinus griseimembra monkeys. J. Parasitol. 75:61-65.[CrossRef][Medline]
    34. 34
  34. Collins, W. E., J. C. Skinner, J. R. Broderson, B. B. Richardson, and P. S. Stanfill. 1989. The Uganda I/CDC strain of Plasmodium malariae in Saimiri sciureus boliviensis. J. Parasitol. 75:310-313.[CrossRef][Medline]
    35. 35
  35. Collins, W. E., J. C. Skinner, and R. E. Coifman. 1967. Fluorescent antibody studies in human malaria. V. Response of sera from Nigerians to five Plasmodium antigens. Am. J. Trop. Med. Hyg. 16:568-571.[Abstract/Free Full Text]
    36. 36
  36. Collins, W. E., J. C. Skinner, M. Pappaioanou, J. R. Broderson, N. S. Ma, V. Filspski, P. S. Stanfill, and L. Rogers. 1988. Infection of Peruvian Aotus nancymai monkeys with different strains of Plasmodium falciparum, P. vivax, and P. malariae. J. Parasitol. 74:392-398.[CrossRef][Medline]
    37. 37
  37. Collins, W. E., J. S. Sullivan, D. Nace, T. Williams, J. Kendall, J. J. Sullivan, G. G. Galland, K. K. Grady, and A. Bounngaseng. 2002. Experimental infection of Anopheles farauti with different species of Plasmodium. J. Parasitol. 88:295-298.[CrossRef][Medline]
    38. 38
  38. Collins, W. E., M. Warren, J. C. Skinner, and H. J. Fredericks. 1968. Studies on the relationship between fluorescent antibody response and ecology of malaria in Malaysia. Bull. W. H. O. 39:451-463.[Medline]
    39. 39
  39. Constantinescu, P., and E. Negulici. 1967. The experimental transmission of Plasmodium malariae to Anopheles labranchiae atroparvus. Trans. R. Soc. Trop. Med. Hyg. 61:182-188.[CrossRef][Medline]
    40. 40
  40. Contacos, P. G., and W. E. Collins. 1969. Plasmodium malariae: transmission from monkey to man by mosquito bite. Science 165:918-919.[Abstract/Free Full Text]
    41. 41
  41. de Arruda, M., E. H. Nardin, R. S. Nussenzweig, and A. H. Cochrane. 1989. Sero-epidemiological studies of malaria in Indian tribes and monkeys of the Amazon Basin of Brazil. Am. J. Trop. Med. Hyg. 41:379-385.[Medline]
    42. 42
  42. de Buck, A. 1935. Infection experiments with quartan malaria. Ann. Trop. Med. Parasit. 29:171-175.
    43. 43
  43. Deloron P., G. H. Campbell, D. Brandling-Bennett, J. M. Roberts, I. K. Schwartz, J. S. Odera, A. A. Lal, C. O. Osanga, V. de la. Cruz, and T. M. McCutchan. 1989. Antibodies to Plasmodium falciparum ring-infected erythrocyte surface antigen and P. falciparum and P. malariae circumsporozoite proteins: seasonal prevalence in Kenyan villages. Am. J. Trop. Med. Hyg. 41:395-399.[Abstract/Free Full Text]
    44. 44
  44. Dixon, F. J. 1966. Comments on immuno-pathology. Mil. Med. 131(Suppl.):1233-1234.[Medline]
    45. 45
  45. Edington, G. M., and H. M. Gilles. 1976. Pathology in the tropics, p. 616-652. Arnold, London, United Kingdom.
    46. 46
  46. Escalante, A. A., D. E. Freeland, W. E. Collins, and A. A. Lal. 1998. The evolution of primate malaria parasites based on the gene encoding cytochrome b from the linear mitochrondrial genome. Proc. Natl. Acad. Sci. USA 95:8124-8129.[Abstract/Free Full Text]
    47. 47
  47. Garnham, P. C. C. 1966. Malaria parasite and other haemosporidia, p. 1114. Blackwell Scientific Publications, Oxford, United Kingdom.
    48. 48
  48. Garnham, P. C. C., R. Lainson, and A. E. Gunders. 1956. Some observations on malaria parasites in a chimpanzee, with particular reference to the persistence of Plasmodium reichenowi and Plasmodium vivax. Ann. Soc. Belge Méd. Trop. 36:811-821.[Medline]
    49. 49
  49. Geiman, Q. M., and W. A. Siddiqui. 1969. Susceptibility of a New World monkey to Plasmodium malariae from man. Am. J. Trop. Med. Hyg. 18:351-354.[Abstract/Free Full Text]
    50. 50
  50. Gendel'man, C. A. 1936. Sull'infezione sperimentale dell' Anopheles nigripes con plasmodi. Trydii Tropitchkogo Inst. Narkomzdrava Assr. Abhhazii 2:70.
    51. 51
  51. Giglioli, G. 1930. Malarial nephritis: epidemiological and clinical notes on malaria. Blackwater fever, albuminuria and nephritis in the interior of British Guiana. Based on seven years' continual observation. Churchill Livingstone, London, United Kingdom.
    52. 52
  52. Gilles, H. M., and R. G. Hendrickse. 1960. Possible aetiological role of Plasmodium malariae in "nephritic syndrome" in Nigerian children. Lancet i:806-807.
    53. 53
  53. Gilles, H. M., and R. G. Hendrickse. 1963. Nephrosis in Nigerian children: role of Plasmodium malariae and effect of antimalarial treatment. Br. Med. J. 2:29-31.[Free Full Text]
    54. 54
  54. Golgi, C. 1886. Malarial infection. Arch. Sci. Med. 10:109-135.
    55. 55
  55. Goman, M., B. Mons, and J. Scaife. 1991. The complete sequence of a Plasmodium malariae SSUrRNA gene and its comparison to other plasmodial SSUrRNA genes. Mol. Biochem. Parasitol. 45:281-288.[CrossRef][Medline]
    56. 56
  56. Gordon, R. M., and T. H. Davey. 1932. P. malariae in Freetown, Sierra Leone. Ann. Trop. Med. Parasitol. 26:65-84.
    57. 57
  57. Grassi, B., A. Bignami, and G. Bastianelli. 1899. Risoconto degli studi fatti sulla malaria durante il mese de Gennaio. Rend. Atti Accad. Lincei 8:100-104.
    58. 58
  58. Grassi, B. and R. Feletti. 1890. Parasites malariques chez les oiseaux. Arch. Ital. Biol. 13:297-300.
    59. 59
  59. Green, R. 1929. Observations on some factors influencing the infectivity of malarial gamete carriers in Malaya to Anopheles maculates. Bull. Inst. Med. Res. Fed. Malay States 5:1-41.
    60. 60
  60. Hendrickse, R. G., and A. Adeniyi. 1979. Quartan malaria nephritic syndrome in children. Kidney Int. 16:64-74.[Medline]
    61. 61
  61. Hendrickse, R. G., A. Adeniyi, G. M. Edington, E. F. Glascow, R. H. R. White, and V. Houba. 1972. Quartan malaria nephritic syndrome: collaborative clinicopathological study in Nigerian children. Lancet i:1143-1148.
    62. 62
  62. Hook, E. W., and S. M. Marra. 1992. Acquired syphilis in adults. N. Engl. J. Med. 326:1060-1069.[Medline]
    63. 63
  63. Hylkema, B. 1920. The development of the parasites of quartan malaria in the Myzomyia ludlowi and their transmission to man. Meded. V.d. Burg. Geneesk. D 6:51.
    64. 64
  64. Iyengar, M. O. T. 1933. Experimental infection of anopheline mosquitoes. Indian J. Med. Res. 20:841-861.
    65. 65
  65. Jamcsó, N. 1921. Experimentelle Untersuchungen über die Malaria-infection des Anopheles und des Menschen beeininfussenden Umstände. Arch. F. Schiffsu. Trop. Hyg. 25:5.
    66. 66
  66. James, S. P. 1931. Some general results of a study of induced malaria in England. Trans. R. Soc. Trop. Med. Hyg. 24:477-525.[CrossRef]
    67. 67
  67. Jeffery, G. M., and R. C. Rendtorff. 1955. Preservation of viable human malaria sporozoites by low-temperature freezing. Exp. Parasitol. 4:445-454.[CrossRef][Medline]
    68. 68
  68. Jongwutiwes, S., C. Putaporntip, T. Iwasaki, T. Sata, and H. Kanbara. 2004. Naturally acquired Plasmodium knowlesi malaria in human, Thailand. Emerg. Infect. Dis. 10:2211-2213.[Medline]
    69. 69
  69. Khaw, O. K., and N. C. Kan. 1934. Some observations on the prevalence of malaria in Nanking and its vicinity. Chinese Med. J. 48:109-123.
    70. 70
  70. Khodukin, N. I., and A. I. Lisova. 1930. Zur Frage über die Möglichkeit von Erkrankungen an Malaria im Winter. Meditz. Muisl' Uzbekitana 1:76.
    71. 71
  71. Kibukamusoke, J. W., and M. S. R. Hutt. 1967. Histological features of the nephritic syndrome of quartan malaria in Uganda. J. Clin. Pathol. 20:117-123.[Abstract/Free Full Text]
    72. 72
  72. Kitchen, S. F. 1949. Quartan malaria, p. 1017-1026. In M. F. Boyd (ed.), Malariology, vol. II. W. B. Saunders Co., Philadelphia, PA.
    73. 73
  73. Knowles, R., and B. C. Basu. 1934. Laboratory studies on the infectivity of Anopheles stephensi. J. Malaria Inst. India 5:1-29.
    74. 74
  74. Lal, A. A., V. F. de la Cruz, G. H. Campbell, P. M. Procell, W. E. Collins, and T. F. McCutchan. 1988. Structure of the circumsporozoite gene from the human malaria Plasmodium malariae. Mol. Biochem. Parasitol. 30:291-294.[CrossRef][Medline]
    75. 75
  75. Laveran, A. 1880. Note sur un nouveau parasite trouvè dans le sang de plusieurs malades atteints de fièvre palustre. Bull. Acad. Med. 9:1235-1236.
    76. 76
  76. Liu, Q., S. Zhu, S. Mizuno, M. Kimura, P. Liu, S. Isomura, X. Wang, and F. Kawamoto. 1998. Sequence variation in the small-subunit rRNA gene of Plasmodium malariae and prevalence of isolates with the variant sequence in Sichuan, China. J. Clin. Microbiol. 36:3378-3381.[Abstract/Free Full Text]
    77. 77
  77. Lupascu, G., P. Constantinescu, E. Negulici, P. C. C. Garnham, R. S. Bray, R. Killick-Kendrick, P. G. Shute, and M. Maryon. 1967. The late primary exoerythrocytic stages of Plasmodium malariae. Trans. R. Soc. Trop. Med. Hyg. 61:482-489.[CrossRef][Medline]
    78. 78
  78. Lupascu, G., P. Constantinescu, E. Negulici, P. G. Shute, and M. E. Maryon. 1968. Parasitologic and clinical investigations on infections with the VS Roumanian strain of Plasmodium malariae transmitted by Anopheles labranchiae atroparvus. Bull. W. H. O. 38:61-67.[Medline]
    79. 79
  79. Mackerras, M. J., and Q. N. Ercole. 1948. Observations on the life-cycle of Plasmodium malariae. Aust. J. Exp. Biol. Med. Sci. 26:515-519.[CrossRef][Medline]
    80. 80
  80. Mackerras, M. J., and F. H. S. Roberts. 1947. Experimental malarial infections in Australasian anophelines. Ann. Trop. Med. Parasitol. 41:329-356.[Medline]
    81. 81
  81. Marotta, G., and G. Sandicchi. 1939. Contributo all'infezione sperimentale della malattia di anofeli con Plasmodium malariae c inoculazione della malattia all'uomo. Riv. Malariol. 18:89-94.
    82. 82
  82. Mayne, B. 1932. Note on experimental infection of Anopheles punctipennis with quartan malaria. Public Health Rep. 47:1771-1773.
    83. 83
  83. McKenzie, F. E., W. E. Collins, and G. M. Jeffery. 2001. Plasmodium malariae blood-stage dynamics. J. Parasitol. 87:626-638.[CrossRef][Medline]
    84. 84
  84. Reference deleted.
    85. 85
  85. McNamara, D. T., L. J. Kasehagen, B. T. Grimberg, J. Cole-Tobian, W. E. Collins, and P. A. Zimmerman. 2006. Diagnosing infection levels of four human malaria parasite species by a polymerase chain reaction/ligase detection reaction fluorescent microsphere-based assay. Am. J. Trop. Med. Hyg. 74:413-421.[Abstract/Free Full Text]
    86. 86
  86. Mer, G. 1933. Observations on the development of Plasmodium malariae Lav. in Anopheles elutus Edw. Ann. Trop. Med. Parasitol. 27:483-488.
    87. 87
  87. Millet, P., W. E. Collins, T. L. Fisk, and P. Nguyen-dinh. 1988. In vitro cultivation of exoerythrocytic stages of the human malaria parasite Plasmodium malariae. Am. J. Trop. Med. Hyg. 38:470-473.[Abstract/Free Full Text]
    88. 88
  88. Muirhead-Thomson, R. C. 1957. Notes on the characters of P. malariae oocysts of possible value in mixed infections. Am. J. Trop. Med. Hyg. 6:980-986.[Medline]
    89. 89
  89. Nace, D., T. Williams, J. S. Sullivan, A. Williams, G. G. Galland, and W. E. Collins. 2004. Susceptibility of Anopheles farauti to infection with different species of Plasmodium. J. Am. Mosquito Control Assoc. 20:272-276.[Medline]
    90. 90
  90. Nagasawa, H., M. Aikawa, P. M. Procell, G. H. Campbell, W. E. Collins, and C. C. Campbell. 1988. Plasmodium malariae: distribution of circumsporozoite protein in midgut oocysts and salivary gland sporozoites. Exp. Parasitol. 66:27-34.[CrossRef][Medline]
    91. 91
  91. Reichenow, E. 1920. Ueber das vorkommen der malariaparasiten des menshen bei den afrikanischen menschenaffen. Zentbl. Bakteriol. Abt. I Orig. 85:207-216.
    92. 92
  92. Rodhain, J. 1940. Les plasmodiums des anthropoids de l'Afrique centrale et leurs relations avec les plasmodiums humains. Ann. Soc. Belge Med. Trop. 20:489-505.
    93. 93
  93. Rodhain, J. 1940. Les plasmodiums des anthropoids de l'Afrique centrale et leurs relations avec les plasmodiums humains. Récepticité de l'homme au Plasmodium malariae. (Plasmodium rodhaini Brumpt) du chimpanzé. C. R. Soc. Biol. 133:276-277.
    94. 94
  94. Rodhain, J. 1941. Les plasmodiums des anthropoids de l'Afrique centrale et leurs relations avec les plasmodiums humains. Bull. Acad. R. Med. Belgique 6:21-60.
    95. 95
  95. Rodhain, J. 1948. Contribution a l'etude des Plasmodiums des anthropoids Africaines. Ann. Soc. Belge Méd. Trop. 28:39-49.
    96. 96
  96. Rodhain, J., and R. Dellaert. 1943. L'infection á Plasmodium malariae du chimpanzé chez l'homme. Etude d'une premiére souche isolée de l'anthropoide Pan satyrus verus. Ann. Soc. Belge. Med. Trop. 23:19-46.
    97. 97
  97. Rougemont, M., M. Van Saanen, R. Sahli, H. P. Hinrikson, J. Bille, and K. Jaton. 2004. Detection of four Plasmodium species in blood from humans by 18S rRNA gene subunit-based and species-specific real-time PCR assays. J. Clin. Microbiol. 42:5636-5643.[Abstract/Free Full Text]
    98. 98
  98. Russell, P. F., and B. N. Mohan. 1939. On experimental malaria infections in certain Anopheles of south-eastern Madras. J. Malaria Inst. India 2:425-431.
    99. 99
  99. Schuurman, C. J., and A. S. B. Huinink. 1929. A malaria problem in Java's South-coast. Meded. Dienst Volksgezondh. Mederl.-Indië 18:120-142.
    100. 100
  100. Schwetz, J. 1933. Sur les parasites malariens (Plasmodium) des singes supérieurs (Anthropoides) africaines. C. R. Soc. Biol. 112:710-711.
    101. 101
  101. Schwetz, J. 1933. Sur une infection malarienne triple d'un chimpanzé. Zentbl. Bakteriol. Parasitenkd. Infektkrankh. Abt. I Orig. 130:105-110.
    102. 102
  102. Schwetz, J. 1934. Sur le paludisme des pygmées. C. R. Soc. Biol. 115:1228-1229.
    103. 103
  103. Shute, P. G. 1951. Mosquito infection in artificially induced malaria. Br. Med. Bull. 8:56-63.[Free Full Text]
    104. 104
  104. Shute, P. G., and M. Maryon. 1951. Studies in the transmission of Plasmodium malariae by Anopheles mosquitoes. Parasitology 41:292-300.[Medline]
    105. 105
  105. Shute, P. G., and M. Maryon. 1952. A study of human malaria oocysts as an aid to species diagnosis. Trans. R. Soc. Trop. Med. Hyg. 46:275-292.[CrossRef][Medline]
    106. 106
  106. Shute, P. G., and M. Maryon. 1955. Transmission of Plasmodium malariae by laboratory-bred Anopheles maculipennis var. atroparvus Meigan. Ann. Trop. Med. Parasitol. 49:451-454.[Medline]
    107. 107
  107. Siddons, L. B. 1944. The experimental transmission of quartan malaria by Anopheles culicifacies Giles. J. Malaria Inst. India 5:361-373.
    108. 108
  108. Singh, B., L. Kim Sung, A. Matusop, A. Radhakrishnan, S. S. Shamsul, J. Cox-Singh, A. Thomas, and D. J. Conway. 2004. A large focus of naturally acquired Plasmodium knowlesi infections in human beings. Lancet 363:1017-1024.[CrossRef][Medline]
    109. 109
  109. Snounou, G., S. Viriyakosol, W. Jarra, S. Thaithong, and K. N. Brown. 1993. Identification of the four human malaria parasite species in field samples by the polymerase chain reaction and detection of a high prevalence of mixed infections. Mol. Biochem. Parasitol. 58:283-292.[CrossRef][Medline]
    110. 110
  110. Soothill, J. F., and R. G. Hendrickse. 1967. Some immunological studies of the nephritic syndrome in Nigerian children. Lancet ii:629-632.
    111. 111
  111. Stephens, J. W. W., and S. R. Christophers. 1902. The relation of species of anopheles to malarial endemicity. Rep. Malaria Commun. R. Soc. Ser. 7:20.
    112. 112
  112. Strickland, C., D. N. Roy, and S. C. Sen Gupta. 1940. Anopheles maculatus and malaria. Trans. R. Soc. Trop. Med. Hyg. 33:639-652.[CrossRef]
    113. 113
  113. Sur, S. N., H. P. Sarkar, and K. M. Banerji. 1932. Plasmochin as a malarial gametocide. Indian Med. Gaz. 67:490-493.
    114. 114
  114. Swellengrebel, N. H., W. Schüffner, and J. M. H. Swellengrebel-de Graaf. 1919. The susceptibility of anophelines to malarial-infections in Netherlands India. Burgerlijk. Geneesk. Dienst. Ned. Indië 3:1-64.
    115. 115
  115. Tahar, R., P. Ringwald, and L. K. Basco. 1998. Heterogeneity in the circumsporozoite protein gene of Plasmodium malariae isolates from sub-Saharan Africa. Mol. Biochem. Parasitol. 92:71-78.[CrossRef][Medline]
    116. 116
  116. Vinetz, J. M., J. Li, T. F. McCutchan, and D. C. Kaslow. 1998. Plasmodium malariae infection in an asymptomatic 74-year-old Greek woman with splenectomy. N. Engl. J. Med. 338:367-371.[Free Full Text]
    117. 117
  117. Watson, M. 1905. Some clinical features of quartan malaria. Indian Med. Gaz. 40:49-52.
    118. 118
  118. Weijer, C. 1999. Another Tuskegee? Am. J. Trop. Med. Hyg. 61:103.[Abstract]
    119. 119
  119. White, R. H. R. 1973. Quartan malaria nephritic syndrome. Nephron ii:147-162.
    120. 120
  120. Young, M. D., and R. W. Burgess. 1947. The transmission of Plasmodium malariae by Anopheles maculipennis freeborni. Am. J. Trop. Med. 27:39-40.
    121. 121
  121. Young, M. D., and R. W. Burgess. 1961. The infectivity to mosquitoes of Plasmodium malariae. Am. J. Hyg. 73:182-192.[Medline]
    122. 122
  122. Young, M. D., N. F. Hardman, R. W. Burgess, W. C. Frohne, and C. W. Sabrosky. 1948. The infectivity of native malarias in South Carolina to Anopheles quadrimaulatus. Am. J. Trop. Med. 28:303-311.[Medline]
    123. 123
  123. Young, M. D., G. R. Coatney, and S. B. McLendon. 1941. Studies on induced quartan malaria in Negro patients. III. Measurements of the paroxysmal phases. South. Med. J. 34:709-712.

Clinical Microbiology Reviews, October 2007, p. 579-592, Vol. 20, No. 4
0893-8512/07/$08.00+0     doi:10.1128/CMR.00027-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.




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