Plasmodium — Details

Plasmodium is a genus of Apicomplexan parasites. Infection by these organisms is known as malaria. The genus Plasmodium was described in 1885 by Ettore Marchiafava and Angelo Celli. Currently over 200 species of this genus are recognized and new species continue to be described.^[1][2]

Of the over 200 known species of Plasmodium, at least 11 species infect humans. Other species infect other animals, including monkeys, rodents, birds, and reptiles. The parasite always has two hosts in its life cycle: a vector—usually a mosquito—and a vertebrate host.

History [edit]

The organism itself was first seen by Laveran on November 6, 1880 at a military hospital in Constantine, Algeria, when he discovered a microgametocyte exflagellating. In 1885, similar organisms were discovered within the blood of birds in Russia. There was brief speculation that birds might be involved in the transmission of malaria; in 1894 Patrick Manson hypothesized that mosquitoes could transmit malaria. This hypothesis was independently confirmed by the Italian physician Giovanni Battista Grassi working in Italy and the British physician Ronald Ross working in India, both in 1898. Ross demonstrated the existence of Plasmodium in the wall of the midgut and salivary glands of a Culex mosquito using bird species as the vertebrate host. For this discovery he won the Nobel Prize in 1902. Grassi showed that human malaria could only be transmitted by Anopheles mosquitoes. It is worth noting, however, that for some species the vector may not be a mosquito.^{[citation needed]}

Biology [edit]

As a protist, the plasmodium is a eukaryote of the phylum Apicomplexa. Unusual characteristics of this organism in comparison to general eukaryotes include the rhoptry, micronemes, and polar rings near the apical end. The plasmodium is known best for the infection it causes, malaria.

The genome of several Plasmodium species—Plasmodium falciparum, Plasmodium knowlesi, Plasmodium vivax, Plasmodium berghei and Plasmodium yoelii—have been sequenced.^[3][4] All these species have genomes of about 25 megabases organised into 14 chromosomes consistent with earlier estimates. The chromosomes vary in length from 500 kilobases to 3.5 megabases and it is presumed that this is the pattern throughout the genus.

The plasmodium contains a degenerated chloroplast called an apicoplast.

The biology of these organisms is more fully described at Plasmodium falciparum biology page.

Diagnostic characteristics of the genus Plasmodium [edit]

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• Merogony occurs both in erythrocytes and other tissues
• Merozoites, schizonts or gametocytes can be seen within erythrocytes
• Merozoites have a "signet-ring" appearance due to a large vacuole that forces the parasite’s nucleus to one pole
• Schizonts are round to oval inclusions that contain the deeply staining merozoites
• Forms gamonts in erythrocytes
• Gametocytes are 'halter-shaped' similar to Haemoproteus but the pigment granules are more confined
• Hemozoin is present
• Vectors are either mosquitos or sandflies
• Vertebrate hosts include mammals, birds and reptiles

Life cycle [edit]

The life cycle of Plasmodium, while complex, is similar to that of several other species in the Haemosporidia.

Vectors [edit]

All the Plasmodium species causing malaria in humans are transmitted by mosquito species of the genus Anopheles. Species of the mosquito genera Aedes, Culex, Culiseta, Mansonia and Theobaldia can also transmit malaria but not to humans. Bird malaria is commonly carried by species belonging to the genus Culex. The life cycle of Plasmodium was discovered by Ross who worked with species from the genus Culex.

Both sexes of mosquitos live on nectar. Because nectar's protein content alone is insufficient for oogenesis (egg production) one or more blood meals is needed by the female. Only female mosquitoes bite.

Biting [edit]

Sporozoites from the saliva of a biting female mosquito are transmitted to skin where they undergo a significant increase in motility in order to traverse skin cells and enter the bloodstream or lymphatic system of the recipient.^[5][6] It has been known for some time now that the parasites block the salivary ducts of the mosquito and as a consequence the insect normally requires multiple attempts to obtain blood. The reason for this has not been clear. It is now known that the multiple attempts by the mosquito may contribute to immunological tolerance of the parasite.^[7] The majority of sporozoites appear to be injected into the subcutaneous tissue from which they migrate into the capillaries and travel to the liver. A proportion are ingested by macrophages and still others are taken up by the lymphatic system where they are presumably destroyed. About 10% of the parasites inoculated by the mosquitoes remain in the skin where they may develop into infective merozoites but do not contribute significantly to systemic infection.^[8]

Dendritic cells [edit]

It is known that the murine parasites can infect, survive and replicate within plasmacytoid dendritic cells of the spleen and that these infections may be productive.^[9][10] The importance of this site of replication in mice has yet to be established and it is currently unknown if these cells support parasite replication in other species.

Hepatic stages [edit]

The majority of sporozoites migrate to the liver and invade hepatocytes. For reasons that are currently unclear each sporozoite typically penetrates several hepatocytes before choosing one to reside within. Once the sporozoite has ceased migration it undergoes an initial remodelling of the pellicle, with disassembly of the inner membrane complex and the appearance of a bulb that progressively enlarges until the initially elongated sporozoite has transformed into a rounded form.^[11][12] This rounded form then matures within the hepatocyte to a schizont containing many merozoites. In some Plasmodium species, such as Plasmodium vivax and Plasmodium ovale, the parasite in the hepatocyte may not achieve maturation to a schizont immediately but remain as a latent or dormant form called a hypnozoite. Although Plasmodium falciparum is not considered to have a hypnozoite form,^[13] this may not be entirely correct (vide infra). This stage may be as short as 48 hours in the rodent parasites and as long as 15 days in P. malariae in humans.

There is considerable variation in the appearance of the blood forms between individuals experimentally inoculated at the same time. Even within a single experimentally individual there may be considerable variation in the maturity of the hepatic forms seen on liver biopsy.

A proportion of the hepatic stages may remain within the liver for considerable time - a form known as hypnozoites. Reactivation of the hypnozoites has been reported for up to 30 years after the initial infection in humans. The factors precipating this reactivation are not known. In the species Plasmodium ovale^[14] and Plasmodium vivax,^[15] but not in Plasmodium malariae,^[16][17] hypnozoites have been shown to occur. It is not yet known if hypnozoite reactivaction occurs with any of the remaining species that infect humans but this is presumed to be the case.

The development from the hepatic stages to the erythrocytic stages has, until very recently, been obscure. Within the hepatocyte the parasites develop into huge multinucleated schizonts within a parasitophorous vacuole.^[18] Thousands of merozoites are formed and released into the host cell cytoplasm by complete disintegration of this parasitophorous vacuole membrane. This is associated with degeneration of the host cell's mitochondria and cessation of protein synthesis which is probably due to the lack of mitochondially produced ATP. This process results in death and detachment of the infected hepatocyte and is followed by the formation of merosomes which may contain hundreds or thousands of merozoites.^[19] The membrane of the merosome is then formed from that of the hepatocyte membrane but the hepatocyte proteins within the membrane are lost. In contrast the membrane of the merozoites is formed by repeated invagination of the parasite's own membrane.^[20] This host derived membrane presumably provides protection from the immune system while the merozoites are transported to the lung. These merosomes lodge in the pulmonary capillaries and slowly disintegrate there over 48–72 hours releasing merozoites.^[21] Erythrocyte invasion is enhanced when blood flow is slow and the cells are tightly packed: both of these conditions are found in the alveolar capillaries.

Infection of the liver may be influenced by the iron regulatory hormone hepcidin^[22] and this may play a role in preventing superinfection despite repeated inoculation.

Erythrocyte stages [edit]

After entering the erythrocyte, the merozoite lose one of their membranes, the apical rings, conoid and the rhopteries. Phagotropy commences and both smooth and granular endoplasmic reticulum becomes prominent. The nucleus may become lobulated.

Within the erythrocytes the merozoite grow first to a ring-shaped form and then to a larger trophozoite form. In the schizont stage, the parasite divides several times to produce new merozoites, which leave the red blood cells and travel within the bloodstream to invade new red blood cells. The parasite feeds by ingesting haemoglobin and other materials from red blood cells and serum. The feeding process damages the erythrocytes. Details of this process have not been studied in species other than Plasmodium falciparum so generalizations may be premature at this time.

Erythrocytes infected by Plasmodium falciparum tend to form clumps - rosettes - and these have been linked to pathology caused by vascular occlusion. This rosette formation may be inhibted by heparin. This agent has been used in the past as part of the treatment of malaria but was abandoned because of an increased risk of haemorrhage. Low molecular weight heparin also disrupts rosette formation and may have a lower risk of bleeding in malaria.^[23] Rosetting has been shown to be due to the binding of the erythrocyte major protein (the var gene product) to the ABO blood group protein.^[24] Blood group A is preferred over group B which in turn is preferred over group O. This has been shown to be due to different fits of blood group protein to the erythrocyte major protein. The binding side on the erythrocyte major protein is opposite to the heparin binding site on the same protein.

Rosette formation has also been reported in Plasmodium vivax.^[25]

Although the ABO blood group is associated with severe malaria this association is lost in pregnancy.^[26]

The regulation of the erythrocyte stages is poorly understood. Perhaps the most striking feature of this process is the synchronous development of millions of parasites. The term the Cinderella syndrome has been coined for the tendency of the parasites to release their merozoites simultaneously during the night.^[27] It is known that melatonin plays a role but how this affects the parasite is only slowly being worked out. It seems that melatonin affects the ubiquitin/proteasome system and a protein kinase (PfPK7) are central to this process.^[28]

The presence of the parasite within the erythrocyte increases the membrane stiffness.^[29] This may be due to an increase in the cross linking of the intraerythrocytic spectrin network or simply due to the mechanical effects of the presence of the parasite itself within the cell.

Merozoites [edit]

The budding of the merozoites from interconnected cytoplasmic masses (pseudocytomeres) is a complex process. At the tip of each bud a thickened region of pellicle gives rise to the apical rings and conoid. As development proceeds an aggregation of smooth membranes and the nucleus enter the base of the bud. The cytoplasm contains numerous large ribosomes. Synchronous multiple cytoplasmic cleavage of the mature schizont results in the formation of numerous uninucleate merozoites.

Escape of the merozoites from the erythrocyte has also been studied.^[30] The erythrocyte swells under osmotic pressure. A pore opens in the erythrocte membrane and 1-2 meorozites escape. This is followed by an eversion the entire erythrocyte membrane, an action that propels the merozoites into the blood stream.

Invasion of erythrocyte precursors has only recently been studied.^[31] The earliest stage susceptible to infection were the orthoblasts - the stage immediately preceding the reticulocyte stage which in turn is the immediate precursor to the mature erythrocyte. Invasion of the erythrocyte is inhibited by angiotensin 2^[32] Angiotensin 2 is normally metabolized by erythrocytes to angiotensin (Ang) IV and Ang-(1-7). Parasite infection decreased the Ang-(1-7) levels and completely abolished Ang IV formation. Ang-(1-7), like its parent molecule, is capable of decreasing the level of infection. The mechanism of inhibition seems likely to be an inhibition of protein kinase A activity within the erythrocyte.

Placental malaria [edit]

More than a hundred late-stage trophozoites or early schizont infected erythrocytes of P. falciparum in a case of placental malaria of a Tanzanian woman were found to form a nidus in an intervillous space of placenta.^[33] While such a concentration of parasites in placental malaria is rare, placental malaria cannot give rise to persistent infection as pregnancy in humans normally lasts only 9 months.

Gametocytes [edit]

Most merozoites continue this replicative cycle but some merozoites differentiate into male or female sexual forms (gametocytes) (also in the blood), which are taken up by the female mosquito. This process of differentiation into gametocytes appears to occur in the bone marrow. Five distinct morphological stages have recognised (stages I - V). Female gametocytes are produced about four times as commonly as male. In chronic infections in humans the gametocytes are often the only forms found in the blood. Incidentally the characteristic form of the female gametocytes in Plasmodium falciparum gave rise to this species's name.

Gameteocytes appear in the blood after a number of days post infection. In P. falciparum infections they appear after 7 to 15 days while in others they appear after 1 to 3 days.^[34] The ratio of asexual to sexual forms is between 10:1 and 156:1^[35][36] The half life of the gametocytes has been estimated to be between 2 and 3 days but some are known to persist for up to four weeks.^[37]

Gametocyte carriage is associated with anaemia.^[38] Although female gametocytes normally outnumber males this may be reversed in the presence of anaemia.

The adhesive properties of the gametocytes have rarely been investigated but they appear to differ from the asexual forms in their adhesive properties.^[39] Stage V gametocytes do not show any appreciable binding, consistent with their condition of being freely circulating cells.

The mechanisms involved in the maturation and release of the gametocytes from the bone marrow are still under investigation. The mature gametocyte infected cells are more deformable than the immature and this is associated with the de association of the STEVOR proteins from the host cell membrane.^[40] It may be that mechanical retention contributes to sequestration of immature gametocytes and that the regained deformability of mature gametocytes is associated with their release in the bloodstream and ability to circulate.

The location of the site of maturation within the bone marrow has rarely been investigated. It appears that the majority stages II-IV may be located with the extravacular spaces within the bone marrow and that only a minority of the mature stages (stage V) are located within this space.^[41]

Gametocyte morphology [edit]

The five recognised morphological stages were first described by Field and Shute in 1956.^[42]

One constant feature of the gametocytes in all stages that distinguishes them from the asexual forms is the presence of a pellicular complex. This originates in small membranous vesicle observed beneath the gametocyte plasmalemma in late stage I. Its function is not known. The structure itself consists of a subpellicular membrane vacuole. Deep to this is an array of longitudinally oriented microtubules. This structure is likely to be relatively inflexible and may help to explain the lack of amoeboid forms observed in asexual parasites.

Gametocyte elongation is driven by the assembly of a system of flattened cisternal membrane compartments underneath the parasite plasma membrane and has a supporting network of microtubules.^[43] The sub-pellicular membrane complex is analogous to the inner membrane complex, an organelle with structural and motor functions that is well conserved across the apicomplexa.

Early stage one gametoctyes are very difficult to distinguish from small round trophozoites. Later stages can be distinguished by the distribution of pigment granulues. Under the electrom microscope the formation of the subpellicular membrane and a smooth plasma membrane are recognisable. The nuclei are recognisably dimorphic into male and female. These forms may be found between day 0 and day 2 in P falciparum infections.

In stage two the gametocyte enlarges and becomes D shaped. The nucleus may occupy a terminal end of the cell or lie along its length. Early spindle formation may be visible. These forms are found between day 1 to day 4 in P falciparum infections.

In stage three the erythrocyte becomes distorted. A staining difference between the male and female gametoctyes is apparent (male stain pink while female stain faint blue with the usual stains). The male nucleus is noticeably larger than the female and more lobulated. The female cytoplasm has more ribosomes, endoplasmic reticulum and mitochondria.

Electron dense organelles (osmophilic bodies) are found in both sexes but are more numerous in the female. The osmophilic bodies are thought to be involved in egress of the gametocyte from the erythrocyte.^[44] These organelles are found between day 4 and day 10 in P. falciparum infections. They are connected to the gametocyte surface by ducts and are almost absent after transformation into the female gamete.

In stage four the erythrocyte is clearly deformed and the gametocyte is elongated. The male gametocytes stain red while the female stain violet blue. In the male pigment granules are scattered while in the female they are more dense. In the male the kinetochores of each chromosome are located over a nuclear pore.

In stage five the gametocytes are clearly recognisable on light microscopy with the typical banana shaped female gametocytes. The subpellicular microtubules depolymerise but the membrane itself remains. In the male gametocyte exhibit the is a dramatic reduction in ribosomal density. Very few mitochondria are retained and the nucleus enlarges with a kinetochore complex attached to the nuclear envelope. In the female gametocytes there are numerous mitochondria, ribosomes and osmophillic bodies. The nucleus is small with a transcription factory.

Stages other than stage five are not normally found in the peripheral blood. For reasons not yet understood stages I to IV are sequestered preferentially in the bone marrow and spleen. Stage V gametocytes only become infectious to mosquitoes after a further two or three days of circulation.

Within the gametocytes are poorly studied Garnham bodies (G bodies).^[45] These are membranous whorls within the cytoplasm and are highly diverse in morphology. They occur in both immature and mature gametocytes. Hemozoin is present within them. Their function is currently unknown.

Infection of mosquito [edit]

The predominant baterium of the Anopheles stephensi is the flavobacterium Elizabethkingia meningoseptica.^[46] What influence this has on the parasite with the typical 24 hour period of its residence within the midgut is not presently known.

In the mosquito's midgut, the gametocytes develop into gametes: the process of activation and gametogenesis occur within 15 minutes of ingestion. and fertilize each other resulting in formation of a diploid zygote: this usually occurs within one hour of ingestion. Zygotes immediately undergo meiosis and differentiate within 24 hours of ingestion into motile, invasive ookinetes. It has been shown that up to 50% of the ookinetes may undergo apoptosis within the midgut.^[47][48] The reason for this behavior is unknown. While in the mosquito gut the parasites form thin cytoplasmic extensions to communicate with each other.^[49] These structures persist from the time of gametocyte activation until the zygote transforms into an ookinete. The function of these tubular structures remains to be discovered.

The ookinetes penetrate the midgut epithelium and escape the midgut, then attach themselves onto the exterior of the gut membrane beneath the basal lamina where they differentiate into oocysts. As in the liver the parasite tends to invade a number of cells before choosing one to reside in. The reason for this behavior is not known. Here they divide many times (usually ~11) to produce large numbers (~8,000) of tiny elongated sporozoites. These sporozoites migrate to the salivary glands of the mosquito where they are injected into the blood and subcutaneous tissue of the next host the mosquito bites.

The pellicle of the ookinete is composed of three membranes: the plasma membrane, and the two linked intermediate and inner membranes which form one flattened vacuole located beneath the plasma membrane.^[50] Beneath this vacuole is found an array of microtubules that are connected to the inner membrane by intramembranous particles. The pellicle differs from all other apicomplexan motile stages by the presence of large pores whose function is currently unknown.

The invasion process appears to be dependent on a serine protease produced by the mosquito in the midgut epithelial cells and in the basal side of the salivary glands.^[51]

The escape of the gametocytes from the erythrocytes has been until recently obscure.^[52] The parasitophorous vacuole membrane ruptures at multiple sites within less than a minute following ingestion. This process may be inhibited by cysteine protease inhibitors. After this rupture of the vacuole the subpellicular membrane begins to disintegrate. This process also can be inhibited by aspartic and the cysteine/serine protease inhibitors. Approximately 15 minutes post-activation, the erythrocyte membrane ruptures at a single breaking point a third process that can be interrupted by protease inhibitors.

Effects on the mosquito

Infection of the mosquito has noticeable effects on the host. The presence of the parasite induces apoptosis of the egg follicles.^[53]

The development of the parasite in the mosquito is temperature dependent with higher temperatures being associated with more rapid development.^[54] Higher temperatures appear to enhance the mosquito's immune system leading to a lower average infection rate.

Survival of infected mosquitoes is enhanced in starvation conditions compared to uninfected controls.^[55] Development within the mosquito involves several insulin like peptides. Blocking this pathway results in reduced parasite development. It appears that the parasite is capable of altering the physiology of the mosquito host and this alternation under starvation conditions is favourable to the host.

Infection appears to reduce fecundity (ability to reproduce) and to increase survival of the mosquito.^[56] This is in line with what evolutionary theory would predict.^{[further explanation needed]}

Ingested peptides within the blood meal may influence the infection rate and survival of the mosquito. Insulin-like growth factor 1 and insulin can persist within the blood meal in the midgut for up to 30 hours. Both peptides can cross the epithelial surface of the midgut and affect the mosquito's physiology. Insulin like growth factor 1 can extended the mosquito lifespan, reduce the rate of infection and the parasite load. Insulin, in contrast, tends to have the opposite effects.^[57]

Mathematical models [edit]

Because of this parasite's importance and complex life cycle models have been developed to help to understand its dynamics.

Life cycle [edit]

The pattern of alternation of sexual and asexual reproduction which may seem confusing at first is a very common pattern in parasitic species. The evolutionary advantages of this type of life cycle were recognised by Gregor Mendel.

Under favourable conditions asexual reproduction is superior to sexual as the parent is well adapted to its environment and its descendents share these genes. Transferring to a new host or in times of stress, sexual reproduction is generally superior as this produces a shuffling of genes which on average at a population level will produce individuals better adapted to the new environment.

The advantages to asexual reproduction within a host can be seen from this simple model taken from Cook.^[58] The proportion of hosts that are parasitised is assumed to be small. This being the case the Poisson distribution is a reasonable model. If the parasite is self-fertilizing then the chance of successful reproduction is 1 - e^-m where m is the proportion of the population parasitised. If the parasite is a faculative bisexual one - one that requires the presence of another parasite on the same host the likelihood of success is 1 - (1 + m)e^-m. If the parasite has two distinct sexes and requires both for reproduction, then the chance of success is ∑ (1 - 2^1-n)(mⁿ / n! e^-m) where the sum is taken between n = 2 and infinity. If m = 0.1 then the chance of success of the self-fertilizing parasite is 40 times that of one with distinct sexes. The chance of success of the bisexual parasite is twice that of the parasite with distinct sexes. For smaller values of m, the advantages of self-fertilization are even greater.

Given that this parasite spends part of its life cycle in two different hosts it must use a proportion of its available resources within each host. The proportion utilized is currently unknown. Empirical estimates of this parameter are desirable for modeling of its life cycle.

Transmission [edit]

The basic model of transmission is known as the Ross-MacDonald model after its authors.^[59][60] It is a set of four coupled non linear ordinary differential equations. This model is also used for other mosquito borne infections including filaria and dengue. While modifications to the model have been proposed to encompass particular conditions,^[61] the basic model is still regarded as a reasonable first approximation to most scenarios.

The model is

In this model X_h is the number of susceptible humans, Y_h is the number of infected humans, X_m is the number of susceptible mosquitoes, Y_m is the number of infected mosquitoes, b is the biting rate, γ is the recovery rate in humans, μ is the mortality rate of the mosquitoes, T_hm is the transmission rate from humans to mosquitoes, T_mh is the transmission rate from mosquitoes to humans and N = X_h + Y_h. The recovery rate is the reciprocal of the mean of the duration of a human infection.

There are a number of simplifying assumptions in this model:

• mosquito and human populations are of constant size
• susceptibility of both humans and mosquitoes to infection is constant
• no incubation period
• neither humans nor mosquitoes are or can become immune to infection
• both populations are well mixed
• the biting rate is constant

Furthermore the equations are deterministic - that is they ignore the possibility of random fluctuations. This model is not solvable in closed form and instead requires numerical integration.

Basic reproductive number [edit]

An important parameter - the basic reproductive number ( R₀ ) - can be derived from this model. This is the average number of new infectious hosts that a typical infectious human will produce during his or her infectious period. An infectious agent can only persist if the basic reproductive number is greater than one. In the Ross-MacDonald model

where m is the number of mosquitoes per human. The term on the right of the equation is the product of the average number of humans infected by a mosquito and the average number of mosquitoes infected by a human. It is worth noting that the biting rate appears here as a quadratic term.

To eliminate an infectious agent it is necessary to reduce the R₀ to below 1. This can be achieved by reducing the biting rate, reducing the number of mosquitoes, shortening the mosquito life span or shortening the duration of infection in the human. If a vaccine is available it is only necessary to vaccinate 1 - 1 / R₀ proportion of the population to eradicate the infectious agent as this will bring R₀ below 1.

This method of estimating R₀ makes a number of simplifying assumptions

• transmission rates are constant
• biting rates are constant
• duration of human infection is constant
• mosquito infection rates are constant

Ignoring the variability that may be present in these parameters can lead to underestimation of R₀.

In the case of malaria in Africa R₀ has been estimated to be 30-815.^[62]

Dormant forms [edit]

Plasmodium falciparum malaria [edit]

A report of P. falciparum malaria in a patient with sickle cell anemia four years after exposure to the parasite has been published.^[63] A second report that P. falciparum malaria had become symptomatic eight years after leaving an endemic area has also been published.^[64]

A third case of an apparent recurrence nine years after leaving an endemic area of P. falciparum malaria has now been reported.^[65] A fourth case of recurrence in a patient with lung cancer has been reported.^[66] Two cases in pregnant women both from Africa but who had not lived there for over a year have been reported.^[67]

A case of congenital malaria due to both P. falciparum and P. malariae has been reported in a child born to a woman from Ghana, a malaria endemic area, despite the mother having emigrated to Austria eighteen months before and never having returned.^[68] A second case of congenital malaria in twins due to P. falciparum has been reported.^[69] The mother had left Togo 14 months before the diagnosis, had not returned in the interim and was never diagnosed with malaria during her pregnancy.

One case of malaria has been reported in a man of African origin with sickle cell trait who was treated for B cell lymphoma with chemotherapy and an autologous bone marrow transplant.^[70] He developed symptomatic malaria only after a subsequent splenectomy performed for worsening disease. Pre treatment blood films and antigen testing were negative.

It seems that at least occasionally P. falciparum has a dormant stage. If this is in fact the case, eradication or control of this organism may be more difficult than previously believed.

Plasmodium malariae [edit]

This parasite is not thought to have a latent form but relapses have been reported.^[71] The mechanism here is not yet clear.

Drug induced [edit]

Developmental arrest was induced by in vitro culture of P. falciparum in the presence of sub lethal concentrations of artemisinin.^[72] The drug induces a subpopulation of ring stages into developmental arrest. At the molecular level this is associated with overexpression of heat shock and erythrocyte binding surface proteins with the reduced expression of a cell-cycle regulator and a DNA biosynthesis protein.

The schizont stage-infected erythrocyte in an experimental culture of P. falciparum, F32 was suppressed to a low level with the use of atovaquone.^[73] The parasites resumed growth several days after the drug was removed from the culture.

Biological refuges [edit]

Macrophages containing merozoites dispersed in their cytoplasm, called 'merophores', were observed in P. vinckei petteri - an organism that causes murine malaria.^[74] Similar merophores were found in the polymorph leukocytes and macrophages of other murine malaria parasite, P. yoelii nigeriensis^[74] and P. chabaudi chabaudi. All these species unlike P. falciparum are known to produce hyponozoites that may cause a relapse. The finding of Landau et al.^[74] on the presence of malaria parasites inside lymphatics suggest a mechanism for the recrudescence and chronicity of malaria infection.^[75]

Starvation induced [edit]

Plasmodium falciparum obtains most of its amino acids from the digestion of haemoglobin. Isoleucine is not present in haemoglobin and must be obtained by alternative methods. When isoleucine is withdrawn from an in vitro culture system, the parasite enters a dormant form.^[76] These dormant forms remain viable for up to 72 hours and resume growth when isoleucine is added to the culture medium. Protein degradation occurs during this time and results in General control nonrepressed 2^{[disambiguation needed]} mediated phosphorylation of eukaryotic initiation factor 2α.

Evolution [edit]

As of 2007^[update], DNA sequences are available from less than sixty species of Plasmodium and most of these are from species infecting either rodent or primate hosts. The evolutionary outline given here should be regarded as speculative, and subject to revision as more data becomes available.

Apicomplexa [edit]

The common ancestor of the Alveolates - a clade to which the Apicomplexa belong - was a myzocytotic predator with two heterodynamic flagella, micropores, trichocysts, rhoptries, micronemes, a polar ring and a coiled open sided conoid.^[77] The Alveolates have lost the axonemal locomotive structures found in the other members of this clade except in gametes.

The ancestor of this group seems likely to have had some photosynthetic ability^[78][79] A recently identified apicomplexan found in Australian corals - Chromera velia - has retained a photosynthetic plastid.^[80] It appears that the alveolates, the dinoflagellates and the heterokont algae acquired their plastids from a red algae suggesting a common origin of this organelle in all these clades.^[81]

Many of the species within the Apicomplexia still possess plastids (the organelle in which photosynthesis occurs in photosynthetic eukaryotes) and some that lack plastids nonetheless have evidence of plastid genes within their genomes. Some extant dinoflagellates can invade the bodies of jellyfish and continue to photosynthesize, which is possible because jellyfish bodies are almost transparent. In host organisms with opaque bodies, such an ability would most likely rapidly be lost. In the majority of such species, the plastids are not capable of photosynthesis. Their function is not known, but there is suggestive evidence that they may be involved in reproduction.

All sequenced mitochondrial genomes of ciliates and apicomplexia are linear.^[82] Whether this is true for the related clades is not yet known. The mitochondrial genome has undergone a severe reduction in size in the Alveolate clade. In the Apicomplexa, where mitochondrion is present, its genome has only three genes (In Cryptosporidium the mitochondion has been lost entirely.) The dinoflagellate mitochondia also have only the same three genes. In Colpodella - a relative of the Apicomplexa - the mitochondrial genome has but a single gene. Since the known ciliate mitochondrial genomes are considerably larger this reduction is genome size must have occurred after their ancestor of this clade diverged from that that gave rise to the extant ciliates. Why this reduction has occurred it not presently clear.

Plasmodium genus [edit]

Current (2007) theory suggests that the genera Plasmodium, Hepatocystis and Haemoproteus evolved from one or more Leucocytozoon species. Parasites of the genus Leucocytozoon infect white blood cells (leukocytes) and liver and spleen cells, and are transmitted by 'black flies' (Simulium species) — a large genus of flies related to the mosquitoes.

Morrison has shown using molecular data that the Haemosporidia are nested within the gregarines.^[83] This clade is a sister to other two clades: the piroplasms and the coccidians

One proposed mechanism of evolution of Leucocytozoon is that its ancestors were parasites spread by the orofaecal route and which infected the intestinal wall. At some point this parasite evolved the ability to infect the liver. This pattern is seen in the genus Cryptosporidium, to which Plasmodium is distantly related. At some later point this ancestor developed the ability to infect blood cells and to survive and infect mosquitoes. Once vector transmission was firmly established, the previous orofecal route of transmission was lost.

The pattern of orofaecal transmission with coincidental infection of erythrocytes is seen in the coccidian genus Schellackia. Species in this genus infect lizards. The usual route of transmission is orofaecal but the parasites can also infect erythrocytes if they traverse the intestinal wall. The infected erythrocytes may be ingested by mites. These infected mites may subsequently be eaten by other uninfected lizards whereupon the parasites emerge and infect these new hosts. Unlike Plasmodium no development occurs in the mite.

In light of Morrison's paper^[83] it would appear that the Haemosporidia evolved from a gregarine that infected mosquitoes. Morrison's data also suggest that the conoid was lost twice independently in all stages with the exception of the merozoite.

Molecular evidence suggests that a reptile - specifically a squamate - was the first vertebrate host of Plasmodium. Birds were the second vertebrate hosts with mammals being the most recent group of vertebrates infected.^[84]

Leukocytes, hepatocytes and most spleen cells actively phagocytose particulate matter, which makes the parasite's entry into the cell easier. The mechanism of entry of Plasmodium species into erythrocytes is still very unclear, as it takes place in less than 30 seconds. It is not yet known if this mechanism evolved before mosquitoes became the main vectors for transmission of Plasmodium.

The genus Plasmodium evolved (presumably from its Leucocytozoon ancestor) about 130 million years ago, a period that is coincidental with the rapid spread of the angiosperms (flowering plants). This expansion in the angiosperms is thought to be due to at least one gene duplication event. It seems probable that the increase in the number of flowers led to an increase in the number of mosquitoes and their contact with vertebrates.

Vectors [edit]

Mosquitoes evolved in what is now South America about 230 million years ago. There are over 3500 species recognized, but to date their evolution has not been well worked out, so a number of gaps in our knowledge of the evolution of Plasmodium remain. There is evidence of a recent expansion of Anopheles gambiae and Anopheles arabiensis populations in the late Pleistocene in Nigeria.^[85]

The reason why a relatively limited number of mosquitoes should be such successful vectors of multiple diseases is not yet known. It has been shown that, among the most common disease-spreading mosquitoes, the symbiont bacterium Wolbachia are not normally present.^[86] It has been shown that infection with Wolbachia can reduce the ability of some viruses and Plasmodium to infect the mosquito, and that this effect is Wolbachia-strain specific.

Classification [edit]

Taxonomy [edit]

Plasmodium belongs to the family Plasmodiidae (Levine, 1988), order Haemosporidia and phylum Apicomplexa. There are currently 450 recognised species in this order. Many species of this order are undergoing reexamination of their taxonomy with DNA analysis.^{[citation needed]} It seems likely that many of these species will be re-assigned after these studies have been completed.^[87][88] For this reason the entire order is outlined here.

Order Haemosporida

Family Haemoproteidae

• Genus Haemocystidium Castellani and Willey 1904, emend. Telford 1996

• Genus Haemoproteus
  • Subgenus Parahaemoproteus
  • Subgenus Haemoproteus

Family Garniidae

• Genus Fallisia Lainson, Landau & Shaw 1974
  • Subgenus Fallisia
  • Subgenus Plasmodioides

• Genus Garnia
• Genus Progarnia

Family Leucocytozoidae

• Genus Leucocytozoon
  • Subgenus Leucocytozoon
  • Subgenus Akiba

Family Plasmodiidae

• Genus Bioccala
• Genus Billbraya
• Genus Dionisia
• Genus Hepatocystis
• Genus Mesnilium
• Genus Nycteria

• Genus Plasmodium
  • Subgenus Asiamoeba Telford 1988
  • Subgenus Bennettinia Valkiūnas 1997
  • Subgenus Carinamoeba Garnham 1966
  • Subgenus Giovannolaia Corradetti, Garnham & Laird 1963
  • Subgenus Haemamoeba Grassi & Feletti 1890
  • Subgenus Huffia Garnham & Laird 1963
  • Subgenus Lacertaemoba Telford 1988
  • Subgenus Laverania Bray 1963
  • Subgenus Novyella Corradetti, Garnham & Laird 1963
  • Subgenus Ophidiella Garnham 1966
  • Subgenus Papernaia Landau et al 2010
  • Subgenus Plasmodium Bray 1963 emend. Garnham 1964
  • Subgenus Paraplasmodium Telford 1988
  • Subgenus Sauramoeba Garnham 1966
  • Subgenus Vinckeia Garnham 1964

• Genus Polychromophilus
• Genus Rayella
• Genus Saurocytozoon
• Genus Vetufebrus Poinar 2011

Phylogenetic trees [edit]

The relationship between a number of these species can be seen on the Tree of Life website. Perhaps the most useful inferences that can be drawn from this phylogenetic tree are:

• P. falciparum and P. reichenowi (subgenus Laverania) branched off early in the evolution of this genus
• The genus Hepatocystis is nested within (paraphytic with) the genus Plasmodium
• The primate (subgenus Plasmodium) and rodent species (subgenus Vinckeia) form distinct groups
• The rodent and primate groups are relatively closely related
• The lizard and bird species are intermingled
• Although Plasmodium gallinaceum (subgenus Haemamoeba) and Plasmodium elongatum (subgenus Huffia) appear be related here there are so few bird species (three) included, this tree may not accurately reflect their real relationship.
• While no snake parasites have been included these are likely to group with the lizard-bird division

While this tree contains a considerable number of species, DNA sequences from many species in this genus have not been included - probably because they are not available yet. Because of this problem, this tree and any conclusions that can be drawn from it should be regarded as provisional.

Three additional trees are available from the American Museum of Natural History.

These trees agree with the Tree of Life. Because of their greater number of species in these trees, some additional inferences can be made:

• The genus Hepatocystis appears to lie within the primate-rodent clade^[89]
• The genus Haemoproteus appears lie within the bird-lizard clade
• The trees are consistent with the proposed origin of Plasmodium from Leucocytozoon

It is also known that the species infecting humans do not form a single clade.^[90] In contrast, the species infecting Old World monkeys seem to form a clade. Plasmodium vivax may have originated in Asia and the related species Plasmodium simium appears to be derived through a transfer from the human P. vivax to New World monkey species in South America. This occurred during an indepth study of Howler Monkeys near São Paulo, Brasil.^[91]

Another tree concentrating on the species infecting the primates is available here: PLOS site

This tree shows that the 'African' (P. malaria and P. ovale) and 'Asian' (P.cynomogli, P. gonderi, P. semiovale and P. simium) species tend to cluster together into separate clades. P. vivax clusters with the 'Asian' species. The rodent species (P. bergei, P. chabaudi and P. yoelli) form a separate clade. As usual P. falciparum does not cluster with any other species. The bird species (P. juxtanucleare, P. gallinaceum and P. relictum) form a clade that is related to the included Leucocytozoon and Haemoproteus species.

A second tree can be found on the PLoS website: PLOS site This tree concentrates largely on the species infecting primates.

The three bird species included in this tree (P. gallinacium, P. juxtanucleare and P. relictum) form a clade.

Four species (P. billbrayi, P. billcollinsi, P. falciparum and P. reichenowi) form a clade within the subgenus Lavernia. This subgenus is more closely related to the other primate species than to the bird species or the included Leuocytozoan species. Both P. billbrayi and P. billcollinsi infect both the chimpanzee subspecies included in this study (Pan troglodytes troglodytes and Pan troglodytes schweinfurthii). P. falciparum infects the bonbo (Pan paniscus) and P. reichenowi infects only one subspecies (Pan troglodytes troglodytes).

The eleven 'Asian' species included here form a clade with P. simium and P. vivax being clearly closely related as are P. knowseli and P. coatneyi; similarly P. brazillium and P. malariae are related. P. hylobati and P. inui are closely related. P. fragile and P. gonderi appear to be more closely related to P. vivax than to P. malariae.

P. coatneyi and P. inui appear to be closely related to P. vivax.^[89]

P. ovale is more closely related to P. malariae than to P. vivax.

Within the 'Asian' clade are three unnamed potential species. One infects each of the two chimpanzee subspecies included in the study (Pan troglodytes troglodytes and Pan troglodytes schweinfurthii). These appear to be related to the P. vivax/P. simium clade.

Two unnamed potential species infect the bonbo (Pan paniscus) and these are related to the P. malariae/P. brazillium clade.

Notes [edit]

An analysis of ten 'Asian' species (P. coatneyi, P. cynomolgi, P. fieldi, P. fragile, P. gonderi, P. hylobati, P. inui, P. knowlesi, P. simiovale and P. vivax) suggests that P. coatneyi and P. knowlesi are closely related and that P. fragile is the species most closely related to these two.^[92] P. vivax and P. cynomolgi appear to be related.

Unlike other eukaryotes studied to date Plasmodium species have two or three distinct SSU rRNA (18S rRNA) molecules encoded within the genome.^[93] These have been divided into types A, S and O. Type A is expressed in the asexual stages; type S in the sexual and type O only in the oocyte. Type O is only known to occur in Plasmodium vivax at present. The reason for this gene duplication is not known but presumably reflects an adaption to the different environments the parasite lives within.

The Asian simian Plasmodium species - Plasmodium coatneyi, Plasmodium cynomolgi, Plasmodium fragile, Plasmodium inui, Plasmodium fieldi, Plasmodium hylobati and Plasmodium simiovale - have a single S-type-like gene and several A-type-like genes. It seems likely that these species form a clade within the subgenus Plasmodium.

Analysis of the merozoite surface protein in ten species of the Asian clade suggest that this group diversified between 3 and 6.3 million years ago - a period that coincided with the radiation of the macques within South East Asia.^[94] The inferred branching order differs from that found from the analysis of other genes suggesting that this phylogenetic tree may be difficult to resolve. Positive selection on this gene was also found.

P. vivax appears to have evolved between 45,000 and 82,000 years ago from a species that infects south east Asian macques.^[95] This is consistent with the other evidence of a south eastern origin of this species.

It has been reported that the C terminal domain of the RNA polymerase 2 in the primate infecting species (other than P. falciparum and probably P. reichenowei) appears to be unusual^[96] suggesting that the classification of species into the subgenus Plasmodium may have an evolutionary and biological basis.

A report of a new species that clusters with P. falciparum and P. reichenowi in chimpanzees has been published, although to date the species has been identified only from the sequence of its mitochondrion.^[97] Further work will be needed to describe this new species, however, it appears to have diverged from the P. falciparum- P. reichenowi clade about 21 million years ago. A second report has confirmed the existence of this species in chimpanzees.^[98] This report has also shown that P. falciparum is not a uniquely human parasite as had been previously believed. A third report of P. falciparum has been published.^[99] This study investigated two mitochondrial genes (cytB and cox1), one plastid gene (tufA), and one nuclear gene (ldh) in 12 chimpanzees and two gorillas from Cameroon and one lemur from Madagascar. Plasmodium falciparum was found in one gorilla and two chimpanzee samples. Two chimpanzee samples tested positive for Plasmodium ovale and one for Plasmodium malariae. Additionally one chimpanzee sample showed the presence of P. reichenowi and another P. gaboni. A new species - Plasmodium malagasi - was provisionally identified in the lemur. This species seems likely to belong to the Vinckeia subgenus but further work is required.

A study of ~3000 wild ape specimens collected from Central Africa has shown that Plasmodium infection is common and is usually with multiple species.^[100] The ape species included in the study were western gorillas (Gorilla gorilla), eastern gorillas (Gorilla beringei), bonobos (Pan paniscus) and chimpanzees (Pan troglodytes). 99% of the strains fell into six species within the subgenus Laverina. P. falciparum formed a monophyletic lineage within the gorilla parasite radiation suggesting an origin in gorrilas rather than chimpanzees.

It has been shown that P. falciparum forms a clade with the species P reichenowi.^[101] This clade may have originated between 3 million and 10000 years ago. It is proposed that the origin of P. falciparum may have occurred when its precursors developed the ability to bind to sialic acid Neu5Ac possibly via erythrocyte binding protein 175. Humans lost the ability to make the sialic acid Neu5Gc from its precursor Neu5Ac several million years ago and this may have protected them against infection with P. reichenowi.

The dates of the evolution of the species within the subgenus Laverania have been estimated as follows:^[102] Laverania: 12.0 million years ago (Mya) (95% estimated range: 6.0 - 19.0 Mya)

P. falciparum in humans: 0.2 Mya (range: 0.078 - 0.33 Mya)

P. falciparum in Pan paniscus: 0.77 Mya (range: 0.43 - 1.6 Mya)

P. falciparum in humans and Pan paniscus: 0.85 Mya (0.46 - 1.3 Mya)

P. reichenowi - P. falciparum in Pan paniscus: 2.2 Mya (range: 1.0 - 3.1 Mya) nd that P. reichenowi - 1.8 Mya (range: 0.6 - 3.2 Mya)

P. billbrayi - 1.1 Mya (range: 0.52 - 1.7 Mya) lciparum P. billcollinsi - 0.97 Mya (range: 0.38 - 1.7 Mya)

Another estimation of the date of evolution of this genus based upon the mutation rate in the cytochrome b gene places the evolution of P. falciparum at 2.5 Mya.^[103] The authors also estimated that the mammalian species of this genus evolved 12.8 Mya and that the order Haemosporida evolved 16.2 Mya. While the date of evolution of P. falciparum is consistent with alternative methods, the other two dates are considerably more recent than other published estimates and probably should be treated with caution.

Plasmodium ovale has recently been shown to consist of two cocirculating species - Plasmodium ovale curtisi and Plasmodium ovale wallikeri.^[104] These two species can only be distinguished by genetic means and they separated between 1.0 and 3.5 million years ago.

A recently (2009) described species (Plasmodium hydrochaeri) that infects capybaras (Hydrochaeris hydrochaeris) may complicate the phylogentics of this genus.^[105] This species appears to be most similar to Plasmodium mexicanum a lizard parasite. Further work in this area seems indicated.

Subgenera [edit]

The full taxonomic name of a species includes the subgenus but this is often omitted. The full name indicates some features of the morphology and type of host species. Sixteen subgenera are currently recognised.

The avian species were discovered soon after the description of P. falciparum and a variety of generic names were created. These were subsequently placed into the genus Plasmodium although some workers continued to use the genera Laverinia and Proteosoma for P. falciparum and the avian species respectively. The 5th and 6th Congresses of Malaria held at Istanbul (1953) and Lisbon (1958) recommended the creation and use of subgenera in this genus. Laverinia was applied to the species infecting humans and Haemamoeba to those infecting lizards and birds. This proposal was not universally accepted. Bray in 1955 proposed a definition for the subgenus Plasmodium and a second for the subgenus Laverinia in 1958. Garnham described a third subgenus - Vinckeia - in 1964.

Mammal infecting species [edit]

Two species in the subgenus Laverania are currently recognised: P. falciparum and P. reichenowi. four additional species - Plasmodium billbrayi, Plasmodium billcollinsi and Plasmodium gaboni - may also exist (based on molecular data) but a full description of these species have not yet been published.^[102][106] The presence of elongated gametocytes in several of the avian subgenera and in Laverania in addition to a number of clinical features suggested that these might be closely related. This is no longer thought to be the case.

The type species is Plasmodium falciparum.

Species infecting monkeys and apes (the higher primates) other than those in the subgenus Laverania are placed in the subgenus Plasmodium. The position of the recently described Plasmodium GorA and Plasmodium GorB has not yet been settled.^[98] The distinction between P. falciparum and P. reichenowi and the other species infecting higher primates was based on the morphological findings but have since been confirmed by DNA analysis.

The type species is Plasmodium malariae.

Parasites infecting other mammals including lower primates (lemurs and others) are classified in the subgenus Vinckeia. Vinckeia while previously considered to be something of a taxonomic 'rag bag' has been recently shown - perhaps rather surprisingly - to form a coherent grouping.

The type species is Plasmodium bubalis.

Bird infecting species [edit]

The remaining groupings are based on the morphology of the parasites. Revisions to this system are likely to occur in the future as more species are subject to analysis of their DNA.

The four subgenera Giovannolaia, Haemamoeba, Huffia and Novyella were created by Corradetti et al. for the known avian malarial species.^[107] A fifth—Bennettinia—was created in 1997 by Valkiunas.^[108] The relationships between the subgenera are the matter of current investigation. Martinsen et al. 's recent (2006) paper outlines what is currently (2007) known.^[109] The subgenera Haemamoeba, Huffia, and Bennettinia appear to be monphylitic. Novyella appears to be well defined with occasional exceptions. The subgenus Giovannolaia needs revision.^[110]

P. juxtanucleare is currently (2007) the only known member of the subgenus Bennettinia.

Nyssorhynchus is an extinct subgenus of Plasmodium. It has one known member - Plasmodium dominicum

Reptile infecting species [edit]

Unlike the mammalian and bird malarias those species (more than 90 currently known) that infect reptiles have been more difficult to classify.

In 1966 Garnham classified those with large schizonts as Sauramoeba, those with small schizonts as Carinamoeba and the single then known species infecting snakes (Plasmodium wenyoni) as Ophidiella.^[111] He was aware of the arbitrariness of this system and that it might not prove to be biologically valid. Telford in 1988 used this scheme as the basis for the currently accepted (2007) system.^[112]

These species have since been divided into 8 genera - Asiamoeba, Carinamoeba, Fallisia, Garnia, Lacertamoeba, Ophidiella, Paraplasmodium and Sauramoeba. Three of these genera (Asiamoeba, Lacertamoeba and Paraplasmodium) were created by Telford in 1988. Another species (Billbraya australis) described in 1990 by Paperna and Landau and is the only known species in this genus. This species may turn out to be another subgenus of lizard infecting Plasmodium.

Classification criteria for subgenera [edit]

Bird infecting species [edit]

This section does not cite any references or sources. Please help improve this section by adding citations to reliable sources. Unsourced material may be challenged and removed. (November 2010)

There are ~40 recognised bird species. Although over 50 species have been described, several have been rejected as being invalid.

With the exception of P. elongatum the exoerythrocytic stages occur in the endothelial cells and those of the macrophage-lymphoid system. The exoerythrocytic stages of P. elongatum parasitise the blood forming cells.

The various subgenera are first distinguished on the basis of the morphology of the mature gametocytes. Those of subgenus Haemamoeba are round or oval while those of the subgenera Giovannolaia, Huffia and Novyella are elongated. These latter genera are distinguished on the basis of the size of the schizonts: Giovannolaia and Huffia have large schizonts while those of Novyella are small.

Species in the subgenus Bennettinia have the following characteristics:

The type species is Plasmodium juxtanucleare.

Species in the subgenus Giovannolaia have the following characteristics:

• Schizonts contain plentiful cytoplasm, are larger than the host cell nucleus and frequently displace it. They are found only in mature erythrocytes.
• Gametocytes are elongated.
• Exoerythrocytic schizogony occurs in the mononuclear phagocyte system.

The type species is Plasmodium circumflexum.

Species in the subgenus Haemamoeba have the following characteristics:

• Mature schizonts are larger than the host cell nucleus and commonly displace it.
• Gametocytes are large, round, oval or irregular in shape and are substantially larger than the host nucleus.

The type species is Plasmodium relictum.

Species in the subgenus Huffia have the following characteristics:

• Mature schizonts, while varying in shape and size, contain plentiful cytoplasm and are commonly found in immature erthryocytes.
• Gametocytes are elongated.

The type species is Plasmodium elongatum.

Species in the subgenus Novyella have the following characteristics:

• Mature schizonts are either smaller than or only slightly larger than the host nucleus. They contain scanty cytoplasm.
• Gametocytes are elongated. Sexual stages in this subgenus resemble those of Haemoproteus.
• Exoerythrocytic schizogony occurs in the mononuclear phagocyte system

The type species is Plasmodium vaughani.

Reptile infecting species [edit]

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All species in these subgenera infect lizards.

Species in the subgenus Asiamoeba have the following characteristics:

Species in the subgenus Carinamoeba have the following characteristics:

• Schizonts normally give rise to less than 8 merozoites
• Schizonts are normally smaller than the host nucleus

The type species is Plasmodium minasense.

Species in the subgenus Fallisia have the following characteristics:

• Non-pigmented asexual and gametocyte forms are found in leukocytes and thrombocytes

Species in the subgenus Garnia have the following characteristics:

• Pigment is absent

Species in the subgenus Lacertaemoba have the following characteristics:

Species in the subgenus Paraplasmodium have the following characteristics:

Species in the subgenus Sauramoeba have the following characteristics:

• Schizonts normally give rise to more than 8 merozoites
• Schizonts are normally larger than the host nucleus
• Non-pigmented gametocytes are typically the only forms found
• Pigmented forms may be found in the leukocytes occasionally

The type species is Plasmodium agamae.

All species in Ophidiella infect snakes

The type species is Plasmodium weyoni.

Notes

• The erythrocytes of both reptiles and birds retain their nucleus, unlike those of mammals. The reason for the loss of the nucleus in mammalian erythocytes remains unknown.

Species listed by subgenera [edit]

The listing given here by subgenus is incomplete. A full listing of the species is available at List of Plasmodium species.

Host range [edit]

Host range among the mammalian orders is non uniform. At least 29 species infect non human primates; rodents outside the tropical parts of Africa are rarely affected; a few species are known to infect bats, porcupines and squirrels; carnivores, insectivores and marsupials are not known to act as hosts.

The listing of host species among the reptiles has rarely been attempted. Ayala in 1978 listed 156 published accounts on 54 valid species and subspecies between 1909 and 1975.^[113] The regional breakdown was Africa: 30 reports on 9 species; Australia, Asia & Oceania: 12 reports on 6 species and 2 subspecies; Americas: 116 reports on 37 species.

Because of the number of species parasited by Plasmodium further discussion has been broken down into following pages:

• Plasmodium species infecting humans and other primates

• Plasmodium species infecting mammals other than primates

• Plasmodium species infecting birds

• Plasmodium species infecting reptiles

Species reclassified into other genera [edit]

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The following species have been assigned to the genus Plasmodium in the past:

• Hepatocystis epomophori
• Hepatocystis kochi
• Hepatocystis limnotragi Van Denberghe 1937
• Hepatocystis pteropi Breinl 1911
• Hepatocystis ratufae Donavan 1920
• Hepatocystis vassali Laveran 1905
• Haemoemba praecox
• Haemoemba rousseleti
• Garnia gonatodi
• Fallisia siamense

Species of dubious validity [edit]

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The following species are currently regarded as questionable validity (nomen dubium).

See also [edit]

• Malaria
• PlasmoDB

References [edit]

Further reading [edit]

Standard reference books for the identification of Plasmodium species

• Laird, Marshall (1998). Avian Malaria in the Asian Tropical Subregion. Santa Clara, CA: Springer-Verlag TELOS. ISBN 981-3083-19-0. 

• Valkiūnas G. (2004). Avian Malaria Parasites and Other Haemosporidia. CRC Press. ISBN 978-0-415-30097-1. 

• Garnham PCC (1966). Malaria Parasites and Other Haemosporidia. Oxford: Blackwell Science Ltd. ISBN 0-632-01770-8. 

This book is the standard reference work on malarial species classification even if it a little dated now. A number of additional species have been described since its publication.

• Hewitt R (1942). Bird Malaria. Baltimore, MD: The Johns Hopkins Press. 

Other useful references

• Shortt HE (1951). "Life-cycle of the mammalian malaria parasite". Br. Med. Bull. 8 (1): 7–9. PMID 14944807. 

• Baldacci P, Ménard R (October 2004). "The elusive malaria sporozoite in the mammalian host". Mol. Microbiol. 54 (2): 298–306. doi:10.1111/j.1365-2958.2004.04275.x. PMID 15469504. 

• Bledsoe GH (December 2005). "Malaria primer for clinicians in the United States". South. Med. J. 98 (12): 1197–204; quiz 1205, 1230. doi:10.1097/01.smj.0000189904.50838.eb. PMID 16440920. 

Red blood cell infected with malaria

The Plasmodium species infecting primates include the parasites causing malaria in humans.

Species of Plasmodium infecting humans [edit]

• Plasmodium falciparum (the cause of malignant tertian malaria)

• Plasmodium vivax (the most frequent cause of benign tertian malaria)

• Plasmodium ovale curtisi (another, less frequent, cause of benign tertian malaria)

• Plasmodium ovale wallikeri (another, less frequent, cause of benign tertian malaria)

• Plasmodium malariae (the cause of benign quartan malaria)

• Plasmodium knowlesi (the cause of severe quotidian malaria in Southeast Asia)

• Plasmodium brasilianum

• Plasmodium cynomolgi

• Plasmodium cynomolgi bastianellii

• Plasmodium inui^[1]

• Plasmodium rhodiani

• Plasmodium schwetzi^[2][3]

• Plasmodium semiovale

• Plasmodium simium

The first six listed here are the most common species that infect humans. With the use of the polymerase chain reaction additional species have been and are still being identified that infect humans.

One possible experimental infection has been reported with Plasmodium eylesi. Fever and low grade parasitemia were apparent at 15 days. The volunteer (Dr Bennett) had previously been infected by Plasmodium cynomolgi and the infection was not transferable to a gibbon (P. eylesi 's natural host) so this cannot be regarded as definitive evidence of its ability to infect humans. A second case has been reported that may have been a case of P. eylesi but the author was not certain of the infecting species.^[4]

A possible infection with Plasmodium tenue has been reported.^[5] This report described a case of malaria in a three year old black girl from Georgia, USA who had never been outside the US. She suffered from both P. falciparum and P. vivax malaria and while forms similar to those described for P. tenue were found in her blood even the author was skeptical about the validity of the diagnosis.

Confusingly Plasmodium tenue was proposed in the same year (1914) for a species found in birds. The human species is now considered to be likely to have been a misdiagnosis and the bird species is described on the Plasmodium tenue page.

Notes [edit]

falciparum

Until recently the only known host of P. falciparum was humans but this species has also been described in gorillas (Gorilla gorilla)^[6] and bonbos^[7] There has been a single report of P. falciparum in a brown howler monkey (Alouatta guariba) and in black howler monkeys (Alouatta caraya)^[8] but until this is confirmed its validity should be considered dubious.

A possible report of P. falciparum in a greater spot-nosed monkeys (Cercopithecus nictitans) has not been confirmed in a large survey.^[9]

A species that clusters with P. falciparum and P. reichenowi has been identified in Gabon, Africa in chimpanzees (Pan troglodytes).^[10] This appears to have diverged from these two species about 21 million years ago. It has only been identified from the sequence of its mitochondrion to date and further work is needed to characterise the species. A second report has confirmed the existence of this species in chimpanzees.^[6] A third report has confirmed the existence of this species.^[11]

Two additional species within the subgenus Laverania have been identified on the basis of DNA sequences alone: Plasmodium billbrayi and Plasmodium billcollinsi.^[6] and bonbos^[7] P. billbrayi was found in two subspecies of chimpanzee (Pan troglodytes troglodytes and Pan troglodytes schweinfurthii). P. billcollinsi was found in only one subspecies of chimpanzee (Pan troglodytes troglodytes). Further work is needed to characterise these species.

malaria

Humans are currently considered to be the only host for P. malariae. However Rodhain and Dellaert in the 1940s showed with transmission studies that P. malariae was present in chimpanzees.^[12][13] The presence of P. malaria in chimpanzees has been reported in Japan suggesting that this species may be able to act as a host.^[14] A second paper has described the presence of P. malaria in wild chimpanzees.^[11] Another paper has reported several cases of P. malariae in brown howler monkey (Alouatta guariba) and black howler monkeys (Alouatta caraya)^[8] It has been shown that splectomised three-striped night monkey (Aotus trivirgatus) can be infected with P. malariae.^[15] The existence of multiple independent reports seem to suggest that the chimpanzee and possibly other species may act as a host to P. malaria at least occasionally.

vivax

P. vivax will infect chimpanzees. Infection tends to be low grade but may be persistent and remain as source of parasites for humans for some time. P. vivax is also known to infect orangutans^[16] and the brown howler monkey (Alouatta guariba clamitans)^[8] P. vivax has been reported from chimpanzees living in the wild.^[11] It has been suggested that vivax infection of the great apes in Africa may act as a reservoir given the prevalence of Duffy antigen negative humans in this area.^[17]

ovale

Like P. vivax, P. ovale has been shown to be transmittable to chimpanzees. P. ovale has an unusual distribution pattern being found in Africa, Myanmar the Philippines and New Guinea. In spite of its admittedly poor transmission to chimpanzees given its discontigous spread, it is suspected that P. ovale may in fact be a zooenosis with an as yet unidentified host. If this is actually the case, the host seems likely to be a primate. A report has been published suggesting that P. ovale may be a natural parasite of chimpanzees^[18] but this needs confirmation. P. ovale has since been described from chimpanzees living in the wild.^[11] This suggests that human infection with this species may as previously suspected be a zooenosis.

It has been recently shown that P. ovale is actually two genetically distinct species that coexist. These species are Plasmodium ovale curtisi and Plasmodium ovale wallikeri.^[19] These two species separated between 1.0 and 3.5 million years ago.

Other species

The remaining species capable of infecting humans all have other primate hosts.

Plasmodium shortii and Plasmodium osmaniae are now considered to be junior synonyms of Plasmodium inui.

It has been proposed that the species P. gora and P. gorb should be renamed P. adleri and P. blacklocki respectively.

Species previously described as infecting humans but no longer recognised as valid [edit]

Taxonomy in parasitology until the advent of DNA based methods has always been a problem and revisions in this area are continuing. A number of synonyms have been given for the species infecting humans that are no longer recognised as valid.^[20] Since perusal of the older literature may be confusing some currently defunct species names are listed here.

P. camerense
P. causiasium
P. golgi
P. immaculatum
P. laverani var. tertium
P. laverani var. quartum
P. malariae var. immaculatum
P. malariae var. incolor
P. malariae var. irregularis
P. malariae var. parva
P. malariae var. quartanae
P. malariae var. quotidianae
P. perniciosum
P. pleurodyniae
P. praecox
P. quartana
P. quotidianum
P. sedecimanae
P. tenue
P. undecimanae
P. vegesio-tertaniae
P. vivax-minuta

Host records [edit]

Most if not all Plasmodium species infect more than one host: the host records shown here should be regarded as incomplete.

• P. billbrayi - chimpanzees (Pan troglodytes troglodytes, Pan troglodytes schweinfurthii)

• P. billcollinsi - chimpanzees (Pan troglodytes troglodytes, Pan troglodytes schweinfurthii)

• P. bouillize - Cercopithecis campbelli

• P. brasilianum - Alouatta fusca, Alouatta palliata, Alouatta seniculus straminea, Alouatta villosa, several night monkey (Aotus) species, Ateles fusciceps, Ateles geoffroyi, Ateles geoffroyi grisescens, Ateles paniscus, Ateles paniscus paniscus, Ateles paniscus chamek, Brachyteles arachnoides, Callicebus moloch ornatus, Callicebus torquatus, Cebus albifrons, Cebus apella, Cebus capucinus, Cebus capucinus capucinus, Cebus capucinus imitator, Chiropotes chiropotes, Lagothrix cana, Lagothrix infumata, Lagothrix lagotricha, Saimiri boliviense, Saimiri sciureus and Saimiri ustus.

• P. bucki - Lemur macaco macaco

• P. cercopitheci - Cercopithecis nictitans

• P. coatneyi - crab eating macque (Macaca fascicularis) and Javanese long-tailed macaque (Macaca irus), silvered leaf monkey (Presbytis cristatus)

• P. coulangesi - Lemur macaco macaco

• P. cynomolgi - bear macaque (Macaca arctoides), Formosan rock macaque (Macaca cyclopis), crab eating macque (Macaca fascicularis), Javanese long-tailed macaque (Macaca irus), Rhesus monkey (Macaca mulatta), southern pig-tailed macaque (Macaca nemestrina), bonnet macaque (Macaca radiata), toque macaque (Macaca sinica), orangutan (Pongo species), silver leaf monkey (Presbytis cristatus) and Hanuman langur (Presbytis entellus)

• P. eylesi - several gibbon (Hylobates) species including Hylobates lar

• P. falciparum - gorillas (Gorilla gorilla), bonbos (Pan paniscus)

• P. fieldi - the crab eating macque (Macaca fascicularis), the Rhesus monkey (Macaca mulatta), the pig-tailed macaque (Macaca nemestrina), the bonnet macaque (Macaca radiata) and the baboon (Papio doguera).

• P. foleyi - Lemur fulvus rufus

• P. fragile - Aotus lemurinus griseimembra, Aotus nancymaae, Aotus vociferans, crab eating macque (Macaca fascicularis), Rhesus monkey (Macaca mulatta), bonnet macaque (Macaca radiata), toque macaque (Macaca sinica) and Saimiri boliviensis

• P. inui - Formosan rock macaque (Macaca cyclopis), crab eating macque (Macaca fascicularis) and Javanese long-tailed macaque (Macaca irus)

• P. gaboni - chimpanzees (Pan troglodytes)^[11]

• P. georgesi - black crested mangabey (Cercocebus albigena)

• P. girardi - Lemur fulvus rufus, Lemur macaco macaco

• P. gonderi - black crested mangabey (Cercocebus albigena), Cercocebus aterrimus, sooty mangabey (Cercocebus atys), Tana River mangabey (Cercocebus galeritus agilus), crab eating macque (Macaca fascicularis) and drills (Mandrillus leucophaeus)

• P. gora - gorillas (Gorilla gorilla)

• P. gorb - gorillas (Gorilla gorilla)

• P. hylobati - several gibbon (Hylobates) species including Hylobates lar and Hylobates moloch

• P. inui - Aotus lemurinus griseimembra, Aotus nancymaae, Celebes black ape (Cynopithecus niger), Assamese macques (Macaca assamensis), crab eating macque (Macaca fascicularis), Rhesus monkey (Macaca mulatta), southern pig-tailed macaque (Macaca nemestrina), bonnet macaque (Macaca radiata), several Presbytis species, Saimiri boliviensis

• P. jefferyi - several gibbon (Hylobates) species

• P. joyeuxi - Cercopithecis callitricus

• P. knowlesi - crab eating macque (Macaca fascicularis), pig-tailed macaque (Macaca nemestrina) and Presbytis malalophus

• P. knowlesi edesoni - Javanese long-tailed macaque (Macaca irus)

• P. lemuris - lemurs (Lemur collaris, Lemur macaco macaco)

• P. malagasi - lemurs

• P. malariae - brown howler monkey (Alouatta guariba clamitans), black howler monkey (Alouatta caraya) chimpanzee (')^[11]

• P. ovale - chimpanzees (Pan)^[11]

• P. percygarnhami - Lemur macaco macaco

• P. petersi - black crested mangabey (Cercocebus albigena)

• P. pitheci - orangutans (Pongo pygmaeus)

• P. reichenowi - chimpanzee (Pan) species^[11] and gorilla (Gorilla) species

• P. rodhaini - chimpanzee (Pan) species and gorilla (Gorilla) species

• P. sandoshami - Malayan flying lemur (Cynocephalus variegatus)

• P. semnopitheci - northern plains gray langur (Semnopithecus entellus)

• P. schwetzi - chimpanzee (Pan) species and gorilla (Gorilla) species

• P. semiovale - toque macaque (Macaca sinica)

• P. shortii - bonnet macaque (Macaca radiata) and toque macaque (Macaca sinica)

• P. silvaticum - orangutans (Pongo pygmaeus)

• P. simium - several howler monkeys (Alouatta) species including the brown howler monkey (Alouatta fusca) and woolly spider monkey (Brachyteles arachnoides)

• P. uilenbergi - Lemur fulvus fulvus

• P. vivax - orangutans (Pongo species), chimpanzees (Pan),^[11] monkeys Saimiri boliviensis,^[21] Aotus lemurinus griseimambra,^[22] the brown howler monkey (Alouatta guariba clamitans) and tamarins (Saguinus mystax and Saguinus fuscicollis)

• P. youngei - white handed gibbon (Hylobates lar)

Primate groups and Plasmodium species [edit]

New World monkeys of the family Cebidae: P. brasilianum and P. simium

Old World monkeys of the Cercopithecidae family: P. coatneyi, P. cynomolgi, P. fieldi, P. fragile, P.gonderi, P. georgesi, P. inui, P. knowlesi, P. petersi, P. shortti and P. simiovale

Gibbons of the Hylobatidae family: P. eylesi, P. hylobati, P. jefferyi and P. youngi

Orangutans (Pongo): P. pitheci and P. silvaticum

Gorillas and chimpanzees: P. billcollini, P. billbrayii, P. falciparum, P. gabonensi, P. gora, P. gorb, P. reichenowi, P. rodhaini and P. schwetzi

Mosquitoes known to transmit human malaria listed by region [edit]

This listing may be incomplete as the taxonomy of this genus is under revision.

North American

• Anopheles (Anopheles) freeborni
• Anopheles (Anopheles) quadrimaculatus
• Anopheles (Nyssorhynchus) albimarus

Central American

• Anopheles (Anopheles) aztecus
• Anopheles (Anopheles) punctimacula
• Anopheles (Anopheles) pseudopunctipennis
• Anopheles (Nyssorhynchus) albimanus
• Anopheles (Nyssorhynchus) albitarsis
• Anopheles (Nyssorhynchus) aquasalis
• Anopheles (Nyssorhynchus) argyritarsis
• Anopheles (Nyssorhynchus) darlingi

South American

• Anopheles (Anopheles) pseudopunctipennis
• Anopheles (Anopheles) punctimacula
• Anopheles (Kerteszia) bellator
• Anopheles (Kerteszia) cruzii
• Anopheles (Kerteszia) neivai
• Anopheles (Nyssorhynchus) albimanus
• Anopheles (Nyssorhynchus) albitarsis
• Anopheles (Nyssorhynchus) aquasalis
• Anopheles (Nyssorhynchus) argyritarsis
• Anopheles (Nyssorhynchus) braziliensis
• Anopheles (Nyssorhynchus) darlingi
• Anopheles (Nyssorhynchus) nuneztovari
• Anopheles (Nyssorhynchus) triannulatus

North Eurasian

• Anopheles (Anopheles) atroparvus
• Anopheles (Anopheles) messeae
• Anopheles (Anopheles) sacharovi
• Anopheles (Anopheles) sinensis
• Anopheles (Cellia) pattoni

Mediterranean

• Anopheles (Anopheles) atroparvus
• Anopheles (Anopheles) claviger
• Anopheles (Anopheles) labranchiae
• Anopheles (Anopheles) messeae
• Anopheles (Anopheles) sacharovi
• Anopheles (Cellia) hispaniola
• Anopheles (Cellia) superpictus

Afro-Arabian

• Anopheles (Cellia) culicifacies
• Anopheles (Cellia) fluviatilis
• Anopheles (Cellia) hispaniola
• Anopheles (Cellia) multicolor
• Anopheles (Cellia) pharoensis
• Anopheles (Cellia) sergentii

Afrotropical

• Anopheles (Cellia) arabiensis
• Anopheles (Cellia) funestus
• Anopheles (Cellia) gambiae
• Anopheles (Cellia) melas
• Anopheles (Cellia) merus
• Anopheles (Cellia) moucheti
• Anopheles (Cellia) nili
• Anopheles (Cellia) pharoensis

Indo-Iranian

• Anopheles (Anopheles) sacharovi
• Anopheles (Cellia) aconitus
• Anopheles (Cellia) annularis
• Anopheles (Cellia) culicifacies
• Anopheles (Cellia) fluviatilis
• Anopheles (Cellia) jeyporiensis
• Anopheles (Cellia) minimus
• Anopheles (Cellia) philippinensis
• Anopheles (Cellia) pulcherrimus
• Anopheles (Cellia) stephensi
• Anopheles (Cellia) sundaicus
• Anopheles (Cellia) superpictus
• Anopheles (Cellia) tessellatus
• Anopheles (Cellia) varuna

Indo-Chinese hills

• Anopheles (Anopheles) nigerrimus
• Anopheles (Cellia) annularis
• Anopheles (Cellia) culicifacies
• Anopheles (Cellia) dirus
• Anopheles (Cellia) fluviatilis
• Anopheles (Cellia) jeyporiensis
• Anopheles (Cellia) maculatus
• Anopheles (Cellia) minimus

Malaysian

• Anopheles (Anopheles) campestris
• Anopheles (Anopheles) conaldi
• Anopheles (Anopheles) donaldi
• Anopheles (Anopheles) letifer
• Anopheles (Anopheles) nigerrimus
• Anopheles (Anopheles) whartoni
• Anopheles (Cellia) acconitus
• Anopheles (Cellia) balabacensis
• Anopheles (Cellia) dirus
• Anopheles (Cellia) flavirostris
• Anopheles (Cellia) jeyporiensis
• Anopheles (Cellia) leucosphyrus
• Anopheles (Cellia) ludlowae
• Anopheles (Cellia) maculatus
• Anopheles (Cellia) mangyanu
• Anopheles (Cellia) minimus
• Anopheles (Cellia) philippiensis
• Anopheles (Cellia) subpictus
• Anopheles (Cellia) sundaicus

Chinese

• Anopheles (Anopheles) anthropophagus
• Anopheles (Anopheles) sinensis
• Anopheles (Cellia) balabacensis
• Anopheles (Cellia) jeyporiensis
• Anopheles (Cellia) pattoni

Australasian

• Anopheles (Anopheles) bacroftii
• Anopheles (Cellia) farauti type 1
• Anopheles (Cellia) farauti type 2
• Anopheles (Cellia) hilli
• Anopheles (Cellia) karwari
• Anopheles (Cellia) koliensis
• Anopheles (Cellia) punctulatus
• Anopheles (Cellia) subpictus

Primate mosquito vectors and associated Plasmodium species [edit]

• Anopheles (Nyssorhynchus) albimanus - P. fieldi, P. vivax

• Anopheles (Nyssorhynchus) albitarsis - P. vivax

• Anopheles (Nyssorhynchus) aquasalis - P. vivax

• Anopheles (Cellia) arabensis - P. falciparum

• Anopheles (Nyssorhynchus) argyritarsi - P. vivax

• Anopheles (Anopheles) argyropus - P. fieldi

• Anopheles (Anopheles) artemievi - P. vivax

• Anopheles (Anopheles) atroparvus - P. fieldi, P. vivax

• Anopheles (Anopheles) aztecus - P. vivax

• Anopheles (Cellia) baimaii - P. vivax

• Anopheles (Cellia) balabacensis - P. fieldi, P. vivax

• Anopheles (Anopheles) beklemishevi - P. vivax

• Anopheles (Kerteszia) bellator - P. vivax

• Anopheles (Nyssorhynchus) benarrochi - P. vivax

• Anopheles (Kertezia) bifurcatus - P. vivax

• Anopheles (Nyssorhynchus) braziliensis - P. vivax

• Anopheles (Kertezia) claviger - P. vivax

• Anopheles (Anopheles) coustani - P. falciparum

• Anopheles (Kerteszia) cruzii - P. vivax

• Anopheles (Cellia) culicifacies - P. vivax

• Anopheles (Nyssorhynchus) darlingi - P. falciparum, P. vivax

• Anopheles (Nyssorhynchus) deaneorum - P. falciparum, P. vivax

• Anopheles (Cellia) dirus - P. cynomolgi, P. fieldi, P. falciparum, P. inui, P. vivax

• Anopheles (Anopheles) donaldi - P. fieldi

• Anopheles (Nyssorhynchus) dunhami - P. vivax

• Anopheles (Cellia) epiroticus - P. vivax

• Anopheles (Cellia) farauti - P. coatneyi, P. vivax^[23]

• Anopheles (Cellia) flavirostris - P. vivax

• Anopheles (Anopheles) freeborni - P. fieldi, P. vivax^[24]

• Anopheles (Cellia) funestus - P. falciparum

• Anopheles (Cellia) fluviatilis - P. vivax

• Anopheles (Cellia) gambiae - P. falciparum, P. vivax

• Anopheles (Cellia) hackeri - P. fieldi, P. knowlesi^[25]

• Anopheles (Kerteszia) homunculus - P. vivax

• Anopheles (Anopheles) hyrcanus - P. vivax

• Anopheles (Cellia) introlatus - P. cynomolgi, P. eylesi

• Anopheles (Anopheles) kleini - P. vivax

• Anopheles (Cellia) kochi - P. eylesi, P. fieldi

• Anopheles (Cellia) latens - P. knowlesi ^[26][27]

• Anopheles (Anopheles) lesteri - P. eylesi

• Anopheles (Anopheles) letifer - P. eylesi, P. fieldi

• Anopheles (Cellia) leucosphyrus - P. eylesi, P. vivax

• Anopheles (Cellia) maculatus - P. eylesi, P. fieldi, P. inui, P. vivax, P. youngei

• Anopheles (Nyssorhynchus) marajoara - P. vivax

• Anopheles (Anopheles) maculipennis - P. vivax

• Anopheles (Anopheles) martinius - P. vivax

• Anopheles (Anopheles) mediopunctatus - P. falciparum, P. vivax

• Anopheles (Cellia) melas - P. falciparum

• Anopheles (Cellia) merus - P. falciparum

• Anopheles (Anopheles) messeae - P. vivax

• Anopheles (Cellia) minimus - P. vivax

• Anopheles (Cellia) moucheti - P. falciparum

• Anopheles (Cellia) nili - P. falciparum

• Anopheles (Nyssorhynchus) nuneztovari - P. vivax

• Anopheles (Nyssorhynchus) oswaldoi - P. falciparum, P. vivax

• Anopheles (Anopheles) paludis - P. falciparum

• Anopheles (Anopheles) peditaeniatus - P. fieldi

• Anopheles (Cellia) philippinensis - P. fieldi

• Anopheles (Anopheles) pseudopunctipennis - P. vivax

• Anopheles (Cellia) pulcherrimus - P. vivax

• Anopheles (Anopheles) pullus - P. vivax

• Anopheles (Anopheles) punctimacula - P. vivax

• Anopheles (Anopheles) punctipennis - P. vivax

• Anopheles (Cellia) quadrimaculatus - P. fieldi, P. vivax

• Anopheles (Nyssorhynchus) rangeli - P. vivax

• Anopheles (Cellia) macarthuri - P. eylesi

• Anopheles (Anopheles) roperi - P. eylesi

• Anopheles (Anopheles) sacharovi - P. vivax

• Anopheles (Cellia) sergentii - P. vivax

• Anopheles sinensis Anopheles (Anopheles) sinensis - P. eylesi, P. fieldi, P. vivax

• Anopheles (Cellia) stephensi - P. cynomogli, P. fieldi, P. inui, P. vivax

• Anopheles (Cellia) sundaicus - P. eylesi, P. vivax, P. youngei

• Anopheles (Cellia) superpictus - P. vivax

• Anopheles (Cellia) tessellatus - P. falciparum, P. vivax

• Anopheles (Nyssorhynchus) triannulatus - P. falciparum, P. vivax

• Anopheles (Nyssorhynchus) trinkae - P. vivax

• Anopheles (Anopheles) umbrosus - P. eylesi

• Anopheles (Cellia) vagus - P. eylesi, P. fieldi

Primate subspecies [edit]

• P. cynomolgi - P. cynomolgi bastianelli, P. cynomolgi ceylonensis and P. cynomolgi cynomolgi.

• P. inui - P. inui inui and P. inui shortii

• P. knowlesi - P. knowlesi edesoni and P. knowlesi knowlesi.

• P. ovale - P. ovale curtisi and P. ovale wallikeri

• P. vivax - P. vivax hibernans, P. vivax chesson and P. vivax multinucleatum.

Interrelatedness - The evolution of these species is still being worked out and the relationships given here should be regarded as tentative. This grouping, while originally made on morphological grounds, now has considerable support at the DNA level.

• P. brasilianum, P. inui and P. rodhaini are similar to P. malariae (quartan malaria group)

• P. cynomolgi, P. fragile, P. knowlesi, P. simium and P. schwetzi are similar to P. vivax

• P. fieldi and P. simiovale are similar to P. ovale

• P. falciparum is closely related to P. reichenowi.

Notes [edit]

• P. kochi has been described as a parasite of monkeys. This species is currently classified as Hepatocystis kochi. This may be subject to revision.

• P. brasilianum and P. rodhaini seem likely to be the same species as P. malariae.

• P. lemuris may actually belong to the Haemoproteus genus. Clarification of this point awaits DNA examination.

• P. shortii is currently (2007) regarded as a junior synonym of P. inui.

References [edit]

Species in six subgenera of Plasmodium infect birds - Bennettinia, Giovannolaia, Haemamoeba, Huffia, Novyella and Papernaia.^[1] Giovannolaia appears to be a polyphytic group and may be sudivided in the future.^[2]

Avian host records

• P. accipiteris - Levant sparrowhawk (Accipiter brevipes)

• P. alloelongatum - Levant sparrowhawk (Accipiter brevipes)

• P. ashfordi - great reed warblers (Acrocephalus arundinaceus),^[3] crossbill (Loxia curvirostra), siskin (Spinus spinus)

• P. bioccai - skylark (Alauda arvensis), magpie (Pica pica)

• P. bigueti - house sparrow (Passer domesticus)^[4]

• P. biziurae - musk duck (Biziura lobata)

• P. buteonis - common buzzard (Buteo buteo)

• P. cathemerium - red-winged blackbird (Agelaius phoeniceus), great horned owl (Bubo virginianus), house finch (Carpodacus mexicanus), blue jay (Cyanocitta cristata), blue tit (Cyanistes caeruleus), wood thrush (Hylocichla mustelina), song sparrow (Melospiza melodia), Northern Mockingbird (Mimus polyglottos leucopterus), cowbirds (Molothrus ater ater), house sparrow (Passer domesticus), magpies (Pica pica budsonia), bronze grackle (Quiscalus quiscula aeneus), northern cardinal (Richmondena cardinalis), canary (Serinus canaria), starling (Sturnus vulgaris), house wren (Troglodytes aedon), robin (Turdus migratorius), white-throated sparrow (Zonotrichia albicollis)

• P. circumflexum - sharp-shinned hawk (Accipiter striatus),^[5] red-winged blackbird (Agelaius phoeniceus), wood duck (Aix sponsa), canvasbacks (Aythya valisineria), blue jay (Cyanocitta cristata), Cape May warbler (Dendroica tigrina), gray cat bird (Dumetella carolinensis), slate colour junicao (Junico hymenalis), song sparrow (Melospiza melodia), common merganser (Mergus merganser), cowbird (Molothrus ater ater), northern cardinal (Richmondena cardinalis cardinalis), trumpeter swan (Olor buccinator), chestnut-tailed starling (Sturnus malabaricus), brown thrasher (Toxostoma rufum), American robin (Turdus migratorius), juniper thrush (Turdus pilaris), wild guineafowl (Numida meleagris)^[6] and white-throated sparrow (Zonotrichia albicollis)

• P. dissanaikei - Ross-ringed parakeet (Psittacula krameri manillensis)

• P. dherteae - skylark (Alauda arvensis), magpie (Pica pica)

• P. dorseti - skylark (Alauda arvensis), magpie (Pica pica)

• P. durae - turkeys (Meleagris species), common peafowl (Pavo cristatus), francolins (Franoclinus leucoscepus and Franoclinus levialanti levialanti), Japanese quail (Coturnix japonica), Lady Amherst pheasants (Chrysophus amherstiae)

• P. elongatum - great reed warblers (Acrocephalus arundinaceus^[7]), red-tailed hawk (Buteo jamaicensis), New Zealand bellbird (Anthornis melanura),^[8] bobwhite quail (Colinus virginianus virginianus), bald eagle (Haliaeetus leucocephalus), honeycreeper (Loxops parva), eastern screech owl (Otus asio), black-footed penguins (Spheniscus demersus),

• P. fallax - pygmy owl (Glaucidium passerinum), turkeys (Meleagris species), helmeted guineafowl (Numida meleagris)

• P. forresteri - eastern screech-owls (Otus asio), great horned owls (Bubo virginianus), barred owls (Strix varia), bald eagles (Haliaeetus leucocephalus), red-shouldered hawks (Buteo lineatus), broad-winged hawks (Buteo platypterus), red-tailed hawks (Buteo jamaicensis)

• P. gabaldoni - muscovy duck (Cairina moschata), rock pigeon (Columba livia)

• P. gallinaceum - red junglefowl (Gallus gallus)

• P. garnhami - rain quail (Coturnix coromandelica)

• P. ghadiriani - skylark (Alauda arvensis), magpie (Pica pica)

• P. giovannolai - red-billed choughs (Pyrrhocorax pyrrhocorax), blackbird (Turdus merula)

• P. globularis - yellow-whiskered greenbul (Andropadus latirostris)

• P. griffithsi - wild turkeys (Meleagris gallopavo intermedia)

• P. gundersi - eastern screech owl (Otus asio)

• P. guangdong - red-whiskered Bulbul (Pycnonotus jocosus)

• P. hegneri - common teal (Anas crecca)

• P. hermani - turkey (Meleagris gallopavo), bobwhite (Colinus virginianus)

• P. hexamerium - skylark (Alauda arvensis), magpie (Pica pica), eastern bluebird (Sialia sialis)

• P. jiangi - red-whiskered bulbul (Pycnonotus jocosus)

• P. juxtanucleare - red junglefowl (Gallus gallus), black-footed penguins (Spheniscus demersus), white eared-pheasant (Crossoptilon crossoptilon)^[9]

• P. kempi - turkey (Meleagris gallopavo), bobwhite (Colinus virginianus), chukar (Alectoris graeca), guinea fowl (Numida meleagris), peacocks (Pavo cristatus), canary (Serinus canaria). Mallards (Anas platyrhynchos) and domestic geese (Anser anser) may be transiently infected.^[10]

• P. loprae - Peking duck (Anas platyrhynchos)

• P. lucens - olive sunbird (Cyanomitra (Nectarinia) olivacea)

• P. lutzi - grey necked wood rail (Aramides cajaneus before: Aramides cajanea) and the great thrush (Turdus fuscater)

• P. matutinum - pigeon and dove (Colubma species)

• P. megaglobularis - olive sunbird (Cyanomitra (Nectarinia) olivacea)

• P. merulae - blackbird (Turdus merula)

• P. mohammedi - house sparrow (Passer domesticus biblicus)

• P. multivacuolaris - yellow-whiskered greenbul (Andropadus latirostris)

• P. nucleophilium - great tit (Parus major), gray catbird (Dumetella carolinensis)

• P. nucleophilum toucani - Swainson's toucan (Ramphastos swainsonii)

• P. octamerium - pintail whydah bird (Vidua macroura)^[11]

• P. pachysomum - tawny pipit (Anthus campestris)

• P. paddae - Java sparrow (Padda oryzivora)

• P. parahexamerium - white-tailed alethe (Alethe diademata)

• P. paranucleophilum - South American tanager

• P. parvulum - vanga species

• P. pedioecetii - lesser prairie-chicken (Tympanuchus pallidicinctus), Darwin's nothura (Nothura darwinii), grouse

• P. pfefferi - magpie (Pica pica)

• P. pinotti - bananaquit (Coereba flaveola), orangequit (Euneornis campestris), yellow-shouldered grassquit (Loxipasser anoxanthus), black-faced grassquit (Tiaris bicolor)

• P. polare - bald eagle (Haliaeetus leucocephalus), barn swallow (Hirundo rustica), yellow wagtails (Motacilla flava)^[12] and cliff swallows (Petrochelidon pyrrhonota

• P. polymorphum - skylark (Alauda arvensis)^[13]

• P. relictum - skylark (Alauda arvensis), reed warbler (Acrocephalus scirpaceus), New Zealand bellbird (Anthornis melanura),^[14] little night owl (Athene noctua), house finch (Carpodacus mexicanus), blue quail (Coturnix chinensis), blue tit (Cyanistes caeruleus), Gyr falcons (Falco rusticolus), chaffinch (Fringilla coelebs), red-backed shrike (Lanius collurio), common crossbill (Loxia curvirostra), Hawaiian honeycreeper, yellow wagtail (Motacilla flava), house sparrow (Passer domesticus), magpie (Pica pica), red-billed choughs (Pyrrhocorax pyrrhocorax), tree sparrow (Passer montanus), great tit (Parus major), the bearded tit (Panurus biarmicus), siskin (Spinus spinus), Magellanic penguins (Spheniscus magellanicus), black-footed penguins (Spheniscus demersus), starling (Sturnus vulgaris), pheasant (Tragopan satyra), white-eyed thrush (Turdus jamaicensis), yellow-faced grassquit (Tiaris olivacea)

• P. rouxi - skylark (Alauda arvensis), partridge

• P. sergentorum - skylark (Alauda arvensis), magpie (Pica pica)

• P. snounoui - magpie (Pica pica)