text:
- Review
- Open Access
Exo-erythrocytic development of avian malaria and related haemosporidian parasites
- Gediminas Valkiūnas1Email author and
- Tatjana A. Iezhova1
© The Author(s) 2017
- Received: 26 January 2017
- Accepted: 20 February 2017
- Published: 3 March 2017
Avian malaria parasites (Plasmodium spp.) and related haemosporidians (Haemosporida) are responsible for diseases which can be severe and even lethal in avian hosts. These parasites cause not only blood pathology, but also damage various organs due to extensive exo-erythrocytic development all over the body, which is not the case during Plasmodium infections in mammals. However, exo-erythrocytic development (tissue merogony or schizogony) remains the most poorly investigated part of life cycle in all groups of wildlife haemosporidian parasites. In spite of remarkable progress in studies of genetic diversity, ecology and evolutionary biology of avian haemosporidians during the past 20 years, there is not much progress in understanding patterns of exo-erythrocytic development in these parasites. The purpose of this review is to overview the main information on exo-erythrocytic development of avian Plasmodium species and related haemosporidian parasites as a baseline for assisting academic and veterinary medicine researchers in morphological identification of these parasites using tissue stages, and to define future research priorities in this field of avian malariology.
The data were considered from peer-reviewed articles and histological material that was accessed in zoological collections in museums of Australia, Europe and the USA. Articles describing tissue stages of avian haemosporidians were included from 1908 to the present. Histological preparations of various organs infected with the exo-erythrocytic stages of different haemosporidian parasites were examined.
In all, 229 published articles were included in this review. Exo-erythrocytic stages of avian Plasmodium, Fallisia, Haemoproteus, Leucocytozoon, and Akiba species were analysed, compared and illustrated. Morphological characters of tissue stages that can be used for diagnostic purposes were specified.
Recent molecular studies combined with histological research show that avian haemosporidians are more virulent than formerly believed. The exo-erythrocytic stages can cause severe disease, especially in non-adapted avian hosts, suggesting the existence of a group of underestimated malignant infections. The development of a given haemosporidian strain can be markedly different in different avian hosts, resulting in significantly different virulence. A methodology combining the traditional histology techniques with molecular diagnostic tools is essential to speed research in this field of avian malariology.
- Avian malaria
- Plasmodium
- Fallisia
- Haemoproteus
- Leucocytozoon
- Akiba
- Exo-erythrocytic development
- Pathology
Malaria parasites (species of Plasmodium) and related haemosporidians of the order Haemosporida are widespread and diverse pathogens infect many species of the major groups of vertebrates, which are exposed to bites of blood-sucking dipteran insects (Diptera). The latter insects are specific vectors of haemosporidians [1]. Many species of reptiles, birds and mammals are often parasitized, and several species of amphibians and even fish have been reported as hosts of haemosporidians [2–6]. The life cycle of these blood parasites includes the following main obligatory stages: (1) exo-erythrocytic merogony (schizogony); (2) development in blood cells and production of gametocytes, which are infective for vectors; and, (3) sexual process and sporogony occurring in dipteran insects, which inject infective sporozoites and initiate new infections in vertebrate hosts. General descriptions of life cycles of different malaria and related haemosporidian species are available in several books and reviews [2–9]. However, the details of development of these pathogens both in avian hosts and vectors remain unknown for the great majority of the described species. This study reviews knowledge and provides comparative morphological data about exo-erythrocytic development of avian malaria and related haemosporidian parasites of the families Plasmodiidae, Haemoproteidae, Garniidae, and Leucocytozoidae.
Avian malaria parasites and other haemosporidians cause not only blood pathology due to development of high parasitaemia, but also pathology in organs because of the damage caused by exo-erythrocytic stages, which often develop in various non-specialized reticuloendothelial cells and can be found all over the body in susceptible birds, including the brain, eyes, nerves, heart, skeletal muscles, and many others organs and tissues [2–7, 10]. From this point of view, avian malaria caused by some Plasmodium species is an even more severe disease than mammalian malaria, during which the exo-erythrocytic development occurs mainly in hepatic cells and usually does not cause disease at this stage of infection [11, 12].
During the last 20 years, numerous studies have addressed the taxonomy, genetic diversity, ecology, evolutionary biology, and genetics of avian haemosporidians (reviewed in [13–19]). The application of sensitive polymerase chain reaction (PCR)-based diagnostic methods, using relatively easy-to-collect samples of infected peripheral blood, provided much new knowledge about these topics, but contributed little information about exo-erythrocytic development. The great majority of recent avian haemosporidian studies only used blood samples to assess haemosporidian infections, which gives no information on the tissue stages. It is important to note that several recent histopathological studies, which combined molecular diagnostic and microscopic research, showed that widespread haemosporidian lineages, particularly of Haemoproteus species, may cause severe disease and even mortality in birds due to marked damage by exo-erythrocytic meronts [20–26]. Interestingly, abortive development of tissue stages of haemosporidians in non-adapted avian hosts caused mortality, and that is a new issue related to bird health in studies of exo-erythrocytic development of avian haemosporidians.
Abortive development happens when a parasite invades a 'wrong' host, succeeds to develop partially, but cannot complete its full life cycle, resulting in the absence of invasive stages (gametocytes in avian hosts or sporozoites in vectors). The parasite is 'captured' by the host during abortive development, which is a dead-end of the infection [27–29]. Infections in 'wrong' hosts are usually eliminated rapidly, with minimal or no symptoms, as was reported in non-successful experimental infections [4, 5]. However, if the infection proceeds partially, the immune response to get rid of it may be highly symptomatic. Abortive haemosporidian infections usually are difficult to diagnose because of unusual location and morphology in the 'wrong' host, unclear etiology, short-term survival of parasites or rapid mortality of infected hosts [22, 28–30].
In old literature (published before 1995), there are numerous descriptions of lethal avian diseases of unclear etiology caused by tissue stages of haemosporidians [31–43]. The tissue stages in dead birds sometimes resemble the megalomeronts of Leucocytozoon species, which is why these diseases were formerly often described as 'aberrant Leucocytozoon infections' or even 'Besnoitia infections' [44]. Recent molecular studies and histopathological research indicate that widespread lineages of haemosporidians are responsible for mortality due to the damage caused by tissue stages of these parasites [20–26, 45]. The application of molecular diagnostic tools indicates that haemosporidian infections might kill abnormal vertebrate hosts during exo-erythrocytic development [22, 29]. For example, the prevalence of Haemoproteus minutus (Haemoproteidae) infection is high (>50%) in most blackbird Turdus merula populations in Europe, and active transmission takes place even in parks of European capitals [26]. The blackbird is adapted to this infection, which is asymptomatic in this host, but H. minutus cause lethal disease in captive parakeets in Europe. Abortive tissue stages of this parasite develop in myocardial and skeletal muscles and other organs, but parasitaemia is absent, making it difficult to diagnose such infections by microscopic examination or PCR-based testing solely of blood samples [22, 23]. Presence of haemosporidians can be reported in organs of birds using solely molecular diagnostic tools [46], but this methodology is insufficiently sensitive to understand pathology caused by the parasites. Traditional histological methods remain important in morphological characterization of haemosporidian tissue stages [47]. However, the descriptions and illustrations of these stages are scattered throughout numerous publications, many of which are old, not easy to access and sometimes are difficult to use for comparative parasitology research.
The aims of this review were: (1) to overview main available information on exo-erythrocytic development of avian Plasmodium and related haemosporidian parasites; and, (2) to discuss future research priorities in this field of avian malariology. This study provides images of common haemosporidian exo-erythrocytic stages, which can be used during identification of these infections. Information about pathological changes in bird organs is only briefly mentioned; it is available in several reviews [2–5, 48] and its detailed consideration requires separate analysis.
Collection of literature
Mainly, full-length papers published in peer-reviewed journals were considered. Old articles (published before 1995) were collected by the first author during his work in the libraries of Oxford University, Natural History Museum, the Liverpool School of Tropical Medicine, and the International Centre for Avian Haematozoa (now available in the Queensland Museum, Australia) in 1992–1996. Published bibliographies of the avian blood-inhabiting haematozoa [49–51] and reviews on haemosporidian parasites [2–5, 7–9, 52, 53] were also used. Recent peer-reviewed articles have been retrieved from online bibliographic databases PubMed, SCOPUS and Google Scholar using the following keywords: 'malaria', 'birds', 'Haemoproteus', 'Plasmodium', 'Leucocytozoon', 'Akiba', 'Fallisia', 'pathology', 'meront', 'schizont'. The Boolean operators 'OR', 'AND', and '>' were used to combine several terms. In all, 229 full text articles and short reports were reviewed. Articles are cited in the review if they explicitly provided data about one of the considered aspects of exo-erythrocytic development or genetic diversity of avian malaria or other haemosporidian parasites; 212 papers containing most representative information about exo-erythrocytic development of these parasites are incorporated in the References.
Collection material and microscopic examination
Histological preparations of exo-erythrocytic stages were obtained from the collections of International Reference Centre for Avian Haematozoa (Queensland Museum, Queensland, Australia), Natural History Museum (London, UK), Nature Research Centre (Vilnius, Lithuania) and the US National Parasite Collection (National Museum of Natural History, Washington DC, USA). All accessed preparations were studied. An Olympus BX61 light microscope (Olympus, Tokyo, Japan) equipped with an Olympus DP70 digital camera and imaging software AnalySIS FIVE (Olympus Soft Imaging Solution GmbH, Münster, Germany) was used to examine preparations and prepare illustrations.
Exo-erythrocytic development of avian malaria parasites
Exoerythrocytic stages reported in different avian Plasmodium parasites
Subgenus and species
Plasmodium cathemerium
P. gallinaceum
P. giovannolai
P. subpraecox
P. lutzi
P. matutinum
P. relictum
P. tejerai
P. circumflexum
P. durae
P. fallax
P. gabaldoni
P. garnhami
P. homocircumflexum
P. lophurae
P. octamerium
P. pinottii
P. polare
P. bertii
P. dissanaikei
P. hexamerium
P. nucleophilum
P. paranucleophilum
P. vaughani
P. juxtanucleare
P. elongatum
P. hermani
P. huffi
The complete exo-erythrocytic development is known only for three species of subgenus Haemamoeba, and three species of subgenus Giovannolaia (Table 1). The tissue stages and the sequence of their development have been relatively well studied in Plasmodium cathemerium, Plasmodium fallax, P. gallinaceum, Plasmodium garnhami, Plasmodium lophurae, and Plasmodium relictum. Experiments with these parasites were relatively easy to design because their vectors (mosquitoes of the genera Culex, Aedes and some others) have been identified and colonized. Additionally, susceptible avian hosts (chickens, ducklings, turkeys, domestic pigeons, and canaries) of these parasites are easy to maintain in the laboratory [2, 4, 5, 52–55, 58, 61, 73, 74, 93, 125]. In these parasites, the general pattern of exo-erythrocytic development is similar, and it is likely to be similar in other species of these subgenera. Mainly, sporozoites induce exo-erythrocytic development in the cells of mesodermal origin, particularly often in the cells of lymphoid-macrophage system and the endothelial cells lining capillaries. Erythrocytic meronts and gametocytes develop in red blood cells. The exo-erythrocytic merogony can be arbitrarily classified in two main stages: the primary (pre-erythrocytic) merogony, which occurs before parasitaemia, and the secondary (post-erythrocytic) merogony, which develops during parasitaemia and maintains at the latency stage. The primary exo-erythrocytic merogony consists at least of two generations of meronts, which are named the cryptozoites and metacryptozoites, respectively. The secondary exo-erythrocytic merogony includes several generations of meronts named the phanerozoites. It is important to note that the development of phanerozoites can be induced by merozoites developed in the erythrocytic meronts, which is not the case in malaria parasites of mammals [2, 4, 5, 11].
Development of the secondary exo-erythrocytic stages (phanerozoites) can be induced in three different ways [10, 54, 55, 61, 64, 65, 67, 69, 73, 82, 91, 92, 94, 97, 117, 127]. First, a proportion of the merozoites from the erythrocytic meronts penetrate the endothelial cells of capillaries and cells of the lymphoid-macrophage system in many organs, including the brain, but particularly often in the spleen and lungs, and produce phanerozoites (Fig. 1e–l). The development of phanerozoites of Plasmodium species can be easily initiated in susceptible birds by merozoites from the erythrocytic meronts, and this feature is often used in experimental research when birds are exposed by the inoculation of infected blood [59, 82, 92, 94, 117, 127]. Second, a proportion of the merozoites developing in the metacryptozoites induce development of phanerozoites. Third, merozoites from phanerozoites can induce further generations of phanerozoites. It is important to note that the secondary exo-erythrocytic merogony (post-erythrocytic merogony or phanerozoite development) is absent in the malaria parasites of mammals, in which only sporozoites initiate exo-erythrocytic development whether directly, or after a delay, when they are responsible for relapses [11].
Maturation of the first generation of phanerozoites often leads to marked increase of parasitaemia. The first phanerozoites usually mature between 10 days and 3 weeks after infection with sporozoites [2, 4, 5, 55, 61, 69, 80, 91]. Phanerozoites develop asynchronously, resulting in the presence of parasites of different maturity in organs of infected birds (Fig. 1h–j). The morphology of phanerozoites is similar to that of metacryptozoites, but the former usually are larger and contain a greater number of merozoites. Over 100 merozoites usually develop in phanerozoites [4].
Both phanerozoites and erythrocytic meronts produce merozoites, which are responsible for maintenance of parasitaemia during the chronic stage of the infection. In addition, phanerozoites are responsible for relapses [2, 55, 69, 70, 91]. Because the merozoites from erythrocytic meronts, metacryptozoites and phanerozoites can initiate secondary exo-erythrocytic merogony, the course of secondary exo-erythrocytic development is complicated and difficult to follow stage by stage. The precise number of generations of phanerozoites is unknown, and it probably varies markedly in different species of parasites and avian hosts.
Both the localization and the sequence of appearance of cryptozoites, metacryptozoites and phanerozoites in different organs and tissues varies markedly depending on mode of infection, parasite species, parasite strain characteristics, age of birds, intensity of infection, stage of infection, and some other factors [4, 10, 53–55, 61, 73, 91, 92, 114, 115, 128].
Exo-erythrocytic meronts (cryptozoites, metacryptozoites, phanerozoites) are covered by a thin wall (Fig. 1a–l). These parasites most often look like roundish or oval bodies containing basophilic cytoplasm and variable numbers of nuclei, which markedly decrease in size as parasites mature (Fig. 1a–e, g–i). The shape of phanerozoites developing in the endothelial cells of capillaries and sinusoids might be determined by the shape of these structures, so they may be elongate, branching or even lobular in shape (Fig. 1j–l); branching and lobular forms particularly occur in brain (Fig. 1j) and muscles. Cytomeres do not usually develop during maturation of exo-erythrocytic meronts of avian Plasmodium parasites.
It is difficult to calculate the number of merozoites in mature exo-erythrocytic meronts because several parasites can develop in one cell or adjacent cells (Fig. 1c), and the boundaries between the parasites are often hardly visible, particularly when parasites are in capillaries (Fig. 1j). The number is very variable however, and fewer than 1000 merozoites usually develop in the exo-erythrocytic meronts of the majority of species of avian malaria parasites. Even young meronts displace the nuclei of infected cells and they can enucleate the host cells, however, they do not usually cause enlargement of nuclei of the host cells (Fig. 1c, g), which is a characteristic feature of exo-erythrocytic development of avian malaria parasites.
Two markedly different types of merozoites have been described in phanerozoites: micro- and macromerozoites (Fig. 1k, l). These have been reported in P. fallax, Plasmodium giovannolai, Plasmodium pinotti and some other species [2, 4, 55, 65, 91, 99]. The micromerozoites predominate in all avian malaria infections; these are small roundish or slightly oval bodies (approximately 1 μm in diameter), each possessing a prominent nucleus and a portion of cytoplasm (Fig. 1f, i, k). Macromerozoites are elongate (between 2 and 5 μm in length), with slightly curved bodies, and one end often slightly rounded; each merozoite possesses a prominent nucleus, and a tiny vacuole was visible in some parasites (Fig. 1l). The role of macromerozoites in the life cycle remains unclear.
The exo-erythrocytic development of the great majority of malaria parasites of subgenera Huffia, Novyella and Bennettinia, remains unknown, or only fragmental data are available, mainly about the development of phanerozoites. The primary exo-erythrocytic meronts (cryptozoites and metacryptozoites) remain undescribed in species belonging to these subgenera because of insufficient experimental research on infections via mosquito bites. Phanerozoites have been reported in some species of these subgenera (Table 1) mainly due to experimental infections, which were induced by inoculation of infected blood.
The exo-erythrocytic merogony of species of subgenus Huffia occurs mainly in cells of the haemopoietic system [55, 56, 116, 117, 120, 123, 124]. Phanerozoites (Fig. 1h, i) are especially numerous in migrating cells of haemopoietic tissues in bone marrow and have also been reported in spleen and liver. They develop in erythroblasts, precursor cells of the erythrocytic series, sometimes in normoblasts and thrombocytes and lymphoid cells, but have also been seen in macrophages, myelocytes and some other related cells. Although some reticuloendothelial cells were seen to be parasitized (for example, macrophages), the fixed reticuloendothelial cells have never been recorded to be parasitized, and phanerozoites have not been seen in the endothelial cells of capillaries in brain or other organs in competent avian hosts, in which the Huffia parasites complete their life cycle and produce gametocytes. However, this pattern of development might change during development in non-competent hosts (see description below). Both the size of phanerozoites and the number of merozoites vary markedly in Huffia species depending on their host cells, but the phanerozoite size rarely exceeds 20 µm in diameter (Fig. 1h, i). Intracellular growing parasites often are surrounded with a light band of the cytoplasm of host cells, and they possess the basophilic cytoplasm and compact bright-stained nuclei (Fig. 1h). Phanerozoites of Huffia species markedly influence the nuclei of host cells (Fig. 1h, i), which is a characteristic feature of the secondary exo-erythrocytic merogony during these infections. The nuclei of infected cells are displaced, deformed and may be even pushed out from the cells (Fig. 1h, i). As a result, some phanerozoites appear to be extracellular bodies (Fig. 1h). Several phanerozoites often parasitize the same host cell during heavy infections.
It is important to note that mode of exo-erythrocytic development can change significantly when Huffia parasites appear in non-adapted avian hosts. For example, enormous numbers of Plasmodium (Huffia) elongatum phanerozoites developed in reticuloendothelial cells in the penguin Spheniscus demersus [129]. In this case, the parasite was found not only in cells of haemopoietic system, but also developed extensively in histiocytes of lungs, spleen, liver, heart, and some other organs. Because of changes in both morphology and site of development of the same parasite during development in different avian hosts, it might be difficult to identify species of Plasmodium using morphological characters of exo-erythrocytic meronts, particularly when infection occurs in unusual hosts.
Available experimental data indicate that the extensive exo-erythrocytic merogony usually lasts about a month post infection. After this period, reticuloendothelial cells can develop resistance to infection, the number of phanerozoites decreases and the parasitaemia maintains mainly due to a limited erythrocytic merogony [2, 4, 61]. It is difficult to see phanerozoites in some Novyella species even if parasitaemia is high [2, 4, 130–132]. Recent histological studies, combined with chromogenic in situ hybridization diagnostic tools indicate that exo-erythrocytic development in some tropical Novyella species is mainly primary, and the persistence of infection is due to long-lasting light parasitaemia [133]. In other words, merozoites developing in erythrocytic meronts of these parasites might be unable or have limited ability to produce phanerozoites. This method of persistence might be common in Novyella species, which are widespread, prevalent and diverse in tropical countries [134, 135], but more investigations are required on their exo-erythrocytic development (Table 1). A similar mode of persistence occurs in Plasmodium malariae in humans [11].
Exo-erythrocytic development of avian garniids
Species of Garniidae are diverse and parasitize many species of reptiles in countries with warm climates. These parasites are similar to species of the Plasmodiidae in that merogony occurs in the cells of fixed tissues and also blood cells of hosts, but malarial pigment (haemozoin) is absent from all stages of their development [138, 139]. Garniids are absent from temperate and cold climates probably due to lack of specific vectors, which remain unidentified [6, 139, 140]. The development of garniids in their vertebrate hosts is similar to that of Plasmodium species in birds, and some species described as garniids belong to this genus [141]. Molecular characterization is needed to specify taxonomic position of haemosporidian species currently belonging to the Garniidae.
Only one species of the Garniidae, Fallisia neotropicalis is known in birds [4, 142]. In this parasite primary exo-erythrocytic meronts, developing from sporozoites have not been reported. Secondary exo-erythrocytic merogony (phanerozoites) can be induced by merozoites developing in the cells of the peripheral blood, as is the case in many species of avian malaria parasites. The phanerozoites look similar to the phanerozoites described in bird malaria parasites. Gabaldon et al. [142] reported that they develop in the reticular cells and in the histiocytes of the connective tissues in many organs including the brain.
It is worth mentioning that there is no DNA sequence information on parasites of the majority of genera and subgenera of the Garniidae, so there is inadequate genetic data to assess their phylogenetic relationships with other haemosporidians. In other words, the entire family is missing from phylogenetic analyses of haemosporidians [15, 17, 143]. That is a shortcoming of the currently available studies on evolution of haemosporidian parasites from the point of view of the limited taxon sampling.
Exo-erythrocytic development of avian haemoproteids
Exoerythrocytic stages reported in different avian Haemoproteus parasites
Subgenus and species
Haemoproteus attenuatus
H. balearicae
H. coatneyi
H. halcionis
H. handai
H. lophortyx
H. mansoni
H. minutus
H. nettionis
H. orizivorae
H. passeris
H. picae
H. sacharovi
H. columbae
H. palumbis
Location of exoerythrocytic stages in different avian haemoproteids
Location of tissue meronts
Parasite subgenus and species
H. columbae, H. palumbis
H. attenuatus, H. balearicae, H. coatneyi, H. nettionis, H. orizivorae, H. passeris, H. picae, H. sacharovi
H. columbae
H. attenuatus, H. coatneyi, H. lophortyx, H. mansoni, H. nettionis, H. picae
H. columbae, H. palumbis
H. coatneyi, H. handai, H. mansoni, H. minutus, H. nettionis
H. columbae
H. coatneyi, H. passeris, H. picae
Skeletal muscles (particularly often in pectoral muscle)
H. columbae
H. halcionis, H. handai, H. mansoni, H. lophortyx
H. columbae
H. coatneyi, H. passeris
H. columbae
H. handai
H. columbae
H. sacharovi
H. coatneyi
H. handai
H. handai
Megalomeronts (Fig. 3c–h) were described in several species of Parahaemoproteus and Haemoproteus (Table 2). These parasites seriously damage affected organs and tissues (Fig. 3e, f), particularly after their maturation and rupture. Megalomeronts of various parasite species have been seen in the skeletal muscle, heart, gizzard, lungs, liver, spleen, kidney, and proventriculus of their hosts [20, 21, 31, 41, 149, 154, 155, 157, 168]. Atkinson et al. [150, 151] reported that skeletal muscles were particularly often parasitized during experimental H. mansoni infection in domestic turkeys (Fig. 3f–h) however, the megalomeronts were also recorded in cardiac muscle. Interestingly, the H. mansoni megalomeronts were not seen in the liver, lungs, brain, kidney, bone marrow, femur, gizzard, duodenum, pancreas, or cecum. Most of these organs, especially lungs and brain, are highly vascular. This suggests a specific requirement for myofibroblasts or their associated physiological environment for the development of megalomeronts. Miltgen et al. [149] also found megalomeronts predominantly in muscle tissues, in the heart, tongue, hip and pectoral muscle; these were especially numerous in the pectoral muscle. Megalomeronts of H. minutus developed in myocardial and skeletal muscles and some other organs of captive parakeets, and the infection was lethal in these avian hosts [23, 26].
Due to their size, the largest megalomeronts can sometimes be seen without a microscope, on the surface of damaged organs [5, 31]. Large parasites are surrounded by a thick hyaline wall (Fig. 3d, f, h). Cytomeres are well defined in developing megalomeronts (Fig. 3d), and nuclei are typically arranged on the periphery of developing cytomeres (Fig. 3i). Megalomeronts are often seen located close to each other, forming large groups (Fig. 3c).
The number of generations of exo-erythrocytic meronts before development of parasitaemia has only been sufficiently documented in H. mansoni, in which at least two generations of the meronts develop in the skeletal and cardiac muscles before the parasite produce merozoites that are able to invade red blood cells [150, 151]. First-generation meronts were seen in the endothelium of capillaries and in myofibroblasts of experimentally infected domestic turkeys. They matured approximately 5 days after infection with sporozoites. The mature parasites can reach up to 20 μm in largest diameter and contain elongate merozoites, which are 5–6 μm in length. The elongate merozoites induce the next generation of merogony in the endothelial cells of the capillaries and in myofibroblasts and also initiate development of meronts in the reticular cells of the spleen. The meronts of the second generation are megalomeronts, which are covered by a hyaline wall (Fig. 3f, h). Megalomeronts mature approximately 17 days after infection and produce numerous roundish merozoites with a diameter of about 1 μm. The merozoites formed in megalomeronts penetrate into red blood cells and become gametocytes.
The overall number of generations of meronts after single-sporozoite infections is unknown in avian haemoproteids, but there are certainly many generations because infected birds usually maintain parasitaemia for many years in controlled laboratory conditions. Additionally, the spring relapses, which are induced by dormant tissue meronts were reported in avian haemoproteids [2, 4, 5]. Available data indicate that meronts of H. mansoni, causing relapses and maintaining chronic parasitaemia in turkeys, develop in reticular cells of the spleen [151].
It is important to note that avian haemoproteids can develop tissue stages in 'wrong' avian hosts, in which sporozoites initiate exo-erythrocytic development, which is then arrested (aborted), so merozoites and gametocytes do not appear. Such abortive infections might be virulent and even lethal in non-adapted avian hosts, but remain insufficiently investigated. Recent PCR-based studies [21–23] have supplemented earlier histopatology research [41, 151] and provided first real evidence that species of Haemoproteus and Parahaemoproteus are responsible for disease and even mortality in birds. It was shown that common species and lineages of Haemoproteus parasites might be relatively benign in naturally adapted hosts, but cause lethal disease in non-adapted birds due to damage of organs by megalomeronts [21–23, 26, 41]. The tissue stages of Haemoproteus species in dead hosts resemble the megalomeronts of Leucocytozoon spp.; it was difficult to determine the disease etiology solely based on morphological data [33–35, 38, 39, 42–44, 169–172]. The traditional opinion about the harmlessness and insignificant veterinary importance of avian haemoproteids [44] requires partial reconsideration [21, 22, 26, 30]. The true extent of pathology and mortality caused by Haemoproteus parasites requires additional investigation, particularly in wildlife because death of infected birds has been reported before the development of parasitaemia. Such disease is difficult to diagnose using blood samples alone, whether either by microscopy or PCR-based tools. Experimental studies are needed, but remain rare because of their difficult design, which requires laboratory manipulations of wild birds and obtaining live sporozoites. In haemoproteids, the latter can mainly be collected by experimental infection of biting midges and hippoboscid flies.
Application of a chromogenic in situ hybridization (ISH) method for detection of tissue meronts in bird organs might be helpful during investigation of exo-erythrocytic merogony of haemosporidians, as is the case in avian Plasmodium parasites [22, 45, 94]. This method speeds up the search for tissue stages of malaria parasites, but has not been used in diagnosis of leucocytozoids and has been rarely applied in avian haemoproteid research [22]. It is important to note that the traditional histology techniques remain informative in haemosporidian parasite morphology research and are useful for better understanding pathologies caused by haemoproteid infections in different organs and tissues (Fig. 3). It worth using these methods in parallel during investigation of exo-erythrocytic development of haemosporidian parasites [94].
Exo-erythrocytic development of avian leucocytozoids
Exoerythrocytic stages reported in different avian Leucocytozoon parasites
Subgenus and species
Leucocytozoon artamidis
L. berestneffi
L. danilewskyi
L. dubreuili
L. fringillinarum
L. lovati
L. macleani
L. marchouxi
L. podargii
L. sakharoffi
L. simondi
L. smithi
L. tawaki
L. caulleryi
During L. dubreuili infection, some of the sporozoites invade liver parenchymal cells and produce small (about 6 μm in largest diameter) oocyst-like meronts, which produced elongate (cystozoite-like) merozoites germinating from a single germinal centre [178]. These meronts are probably persistent (dormant) stages, which are responsible for relapses; they have not been reported in other Leucocytozoon parasites.
After rupture of the hepatic meronts, the cytoplasmic 'fragments' develop in parallel with merozoites. These fragments are called the syncytia (islands); they are covered by a plasma membrane and contain several nuclei [166, 173, 198]. Syncytia are remnants of the meronts. They are unable to penetrate into host cells, but are washed out into the circulation, spread over the bird body and are phagocytized by macrophages and other cells of the reticuloendothelial system giving rise to large meronts—megalomeronts (megaloschizonts) (Fig. 4c–h). The term megalomeront (megaloschizont) was suggested by Huff [162] for the 'host cell-Leucocytozoon parasite' complex. The markedly enlarged nucleus of the host-cell ('central body' of megalomeront) is the most readily distinguishable character of megalomeronts in Leucocytozoon species (Fig. 4c–h). The central body is often located close to the centre of megalomeronts (Fig. 4d, g, h). Syncytia grow rapidly and induce marked enlargement both of the nuclei and the cytoplasm of host cells. Numerous cytomeres appear (Fig. 4d–f). As the parasite matures, each cytomere breaks up into numerous smaller bladder-like portions (subcytomeres, Fig. 4d, e, g), with the nuclei often located on the surface of the subcytomeres. Subdivision of smaller cytomeres continues until uninuclear merozoites appear. Both the division of parasite into cytomeres and the peripheral arrangement of nuclei in the cytomeres facilitate the metabolic and trophic functions. Nuclei of host cells, which contain mature megalomeront, might reach 50–100 μm in diameter and even greater (Fig. 4d, g). The greatly enlarged nucleus of the host cell (the 'central body' of megalomeront) probably takes part in transportation of nutrient materials inside the parasites. Growth of megalomeronts from their initiation to maturity represents a several 1000-fold increase in volume. Mature megalomeronts are packed with numerous merozoites, and boundaries between the cytomeres gradually disappear.
Megalomeronts are surrounded with a thick capsule-like wall of fibrillar structure, which is collagen-positive [166, 173, 195, 198]. The wall can be about 1 µm in width or even thicker. The encapsulation is most obvious in organs with a dense reticular network such as the spleen and lymph nodes (Fig. 4d, e), and it is least in the brain and heart (Fig. 4f). Growing megalomeronts displace surrounding cells and reticular fibres, which become associated with the surface of the expanding sphere and thus take part in the formation of the capsular wall.
Mature megalomeronts are very variable in size, which usually ranges between 50 and 500 µm, but might be even greater. The largest megalomeronts can be seen with the naked eye on the surface of heavily infected organs [4, 166, 173, 195]. It is likely that size of mature megalomeronts partially depends to the size and number of syncytia, which are phagocytized by host cells [166, 195, 199, 202]. Development of megalomeronts is asynchronous due to asynchronous phagocytosis of syncytia in different organs (Fig. 4c).
Megalomeronts have been reported all over body in the infected birds, including brain, nerves, heart, pericardium, lungs, trachea, kidneys, liver, pancreas, intestine, skeletal muscles, masticatory, and glandular parts of stomach, gonads, bursa of Fabricius, but most often seen in the spleen (Fig. 4c–e) and the lymph nodes [162, 166, 173, 186, 191, 192, 195, 205]. Intensity of infection in the latter two organs can be extremely high (Fig. 4c), and the parasites can represent over 50% of the infected spleen mass. The marked enlargement of spleen is sometimes due to the megalomeronts mass per se during intense Leucocytozoon infection [195], and such mechanism of spleen enlargement has not been recorded in any other haemosporidiosis. It should be noted that megalomeronts are usually uncommon in the liver, which is the site of extensive phagocytosis in birds. It seems that the absence or rarity of megalomeronts in the liver is due to local immunity induced after intense primary hepatic merogony (Fig. 4a), and the destruction of syncytia in the Kupffer cells.
Megalomeronts develop rapidly and become mature approximately four to five days after ingestion of syncytia by their host cells or approximately 7–9 days following a sporozoite infection [166, 195, 200, 202]. Many thousands of uninuclear merozoites develop in each megalomeront. The merozoites are approximately of 1 μm in diameter; they inhabit various blood cells and produce gametocytes.
Some of the merozoites formed in megalomeronts are phagocytized by the reticuloendothelial cells and slow down their development, periodically producing megalomeronts, which are responsible for the chronic parasitaemia and also spring relapses [208, 209]. Persisting meronts are few and usually difficult to find. They were seen in lungs during L. simondi infection [199].
Small meronts develop in all species of leucocytozoids, but megalomeronts do not. The latter were reported in L. danilewskyi, Leucocytozoon sakharoffi, L. simondi, L. caulleryi and in some other species (Table 4). Megalomeronts were not found in L. dubreuili, L. fringillinarum, L. smithi and many other species. The role of megalomeronts in the life cycle of leucocytozoids is insufficiently understood. In L. simondi and L. danilewskyi, the merozoites from megalomeronts inhabit blood cells and initiate the development of gametocytes, which markedly change the host cells by producing elongate host cell processes [177, 228]. However, there is no strict correlation between the development of megalomeronts and the appearance of gametocytes in the fusiform host cells in other leucocytozoid species. For example, gametocytes do not develop in fusiform host cells in Leucocytozoon artamidis, L. caulleryi, Leucocytozoon marchouxi, Leucocytozoon podargii and L. sakharoffi, although these species do produce megalomeronts in some avian hosts [174, 186, 187, 191, 192, 226], while such gametocytes predominate in L. smithi, in which megalomeronts are absent [211]. Additional investigations are needed to understand the role of megalomeronts in the life cycle of leucocytozoids.
Exo-erythrocytic merogony of L. (Akiba) caulleryi differs from the species of subgenus Leucocytozoon in several characters [216, 218, 221, 224, 226]. First, the meronts of the first and following generations do not develop in hepatic cells. This feature is more similar to species of Plasmodium and Haemoproteus than to species of Leucocytozoon. All generations of meronts of L. caulleryi grow in the endothelial cells of the capillaries of many organs and can be found in brain, bursa of Fabricius, thymus, trachea, bronchus, lungs, heart, liver, spleen, kidneys, pancreas, gullet, masticatory and glandular stomach, crop, duodenum, ovary, testis, oviduct, and various skeletal muscles. It is interesting to note that they occur not only in the visceral organs but also in the eyes and sciatic nerves [219]. The meronts of L. caulleryi were especially numerous and often seen in bursa of Fabricius and thymus.
Second, merozoites developing in meronts of the first generation are elongate (up to 7 μm in length) and are similar morphologically to the first generation merozoites described in H. mansoni [5, 223, 226].
Main morphological characters of exoerythrocytic meronts of avian haemosporidian parasites
Merogony in hepatocytes (Fig. 4a, b)
Merogony in cells of hemopoietic system (Fig. 1h, i)
Thin-walled elongate meronts in brain capillaries (Fig. 1j–l)
'Central body' in megalomeronts (Fig. 4d–h)
Extracellular development of megalomeronts (Fig. 5a, b)
Prominent (more than threefold in comparison to controls) enlargement of host cell nuclei (Fig. 4d, f–h)
This review shows that the exo-erythrocytic development of the majority of species of wildlife haemosporidian parasites remains unknown or studied only fragmentally (Tables 1, 2 and 4), most often using incidentally collected samples from wild, naturally infected birds, in which the fate and duration of these infections is unclear. Molecular diagnostic tools have opened new opportunities to detect infections and to determine parasite species identity in tissue stages, providing the first real evidence that these avian parasites are more virulent than formerly believed, causing severe damage in various organs of non-adapted avian hosts [21, 22, 26, 29, 30, 94].
It is important to note that the exo-erythrocytic merogony of a given strain of haemosporidian parasites may differ significantly in different avian hosts, sometimes resulting in abortive development and increase of virulence of parasites in 'wrong' hosts. This phenomenon is well documented, but its mechanisms remain insufficiently understood. For example, extensive exo-erythrocytic development of P. elongatum in non-adapted penguins occurs not only in cells of haemopoietic system, as usually is the case in adapted passerines, but also in reticuloendothelial cells, resulting in the intense merogony, the marked damage of various organs and severe disease, during which parasitaemia might be absent [4, 129]. Similar abortive exo-erythrocytic development occurs in H. minutus infection of captive parakeets, in which sporozoites produce megalomeronts that cause lethal disease, but parasitaemia is absent as well [23, 26]. The lethal aberrant avian haemosporidioses caused by tissue stages have been reported in various bird species, using of PCR-based diagnostic methods in Europe, America and Australia [20–26]. Available molecular data leave no doubt that aberrant haemosporidian infections may kill non-adapted avian hosts. However, it is unclear how often that happens in wildlife and what are the consequences of such infections for bird populations.
Many haemosporidian species and their lineages have broad specificity and can infect birds belonging to different species, genera, families, and even orders [4, 5, 13, 14, 16, 19]. The exo-erythrocytic development of a given species of avian haemosporidian can be different in different avian hosts. In other words, the sequence of different stages and their occurrence during the exo-erythrocytic development of a particular parasite strain are not stable characteristics, but they can be functions of the host species. For example, experimental studies showed marked differences in the exo-erythrocytic development of the same strain of Leucocytozoon in different vertebrate hosts [200, 201]. Mainly, a strain of L. simondi developed megalomeronts in ducks, but not in geese. Complete exo-erythrocytic development occurred only ducks. In both species of avian hosts, the infective for vectors stages (gametocytes) developed. The 'partial' exo-erythrocytic development in geese was accompanied by reduced parasitaemia and virulence of this parasite strain. Moreover, the parasite was finally eliminated from infected geese, resulting in lack of relapses, which was not the case in ducks, in which long-lasting parasitaemia and relapses occurred. It seems that ducks are less adapted to some strains of L. simondi, resulting in development of megalomeronts and heavy disease. During development in geese, the same strain undergoes only the 'partial' development, and megalomeronts do not occur. This finding was supported experimentally by exposure of ducks and geese to natural infection of L. simondi at different localities in the upper peninsula of Michigan [200]. Interestingly, L. sakharoffi is cosmopolitan and infects many species of the Corvidae, but megalomeronts were reported only in the hooded crow Corvus cornix [4, 192]. Additionally, Leucocytozoon marchouxi is common in doves and pigeons, but its megalomeronts were reported only in the Mauritian pink pigeon Columba mayeri [186]. It seems that these parasites also develop megalomeronts only in non-adapted avian hosts. Thus, the available data show that the exo-erythrocytic development of haemosporidian parasites is relatively flexible, and it can be modified when the same parasite strain appears in different species of avian host. However, it remains unclear how often such changes occur in wildlife. This issue is related to severity of disease and bird health, and it needs additional investigation. It is essential to expand the application of chromogenic in situ hybridization in research of haemosporidians from simply diagnostic purposes [22, 45, 47, 94] towards species-level detection of localization of the exo-erythrocytic stages and pathology associated with these stages in various tissues. Sampling of naturally dead birds in zoos, aviaries and wildlife as well as experimental infections of captive birds are most straightforward ways to for such research [20–25, 29, 94].
The exo-erythrocytic development of avian haemosporidian parasites requires much additional research. Importantly, the tissue merogony of some avian haemosporidian species and their strains is not as stable as was formerly believed, and it might differ significantly in different avian host species. Understanding the mechanisms of this phenomenon is an important question for future research. Additionally, the use of molecular diagnostic tools and histological methods in parallel have allowed recognition of the existence of underestimated manifestation of haemosporidian infections, which cause diseases and even mortality in birds due to damage of organs by tissue stages. In the latter case, the parasites initiate the exo-erythrocytic development, but cannot complete it in 'wrong' hosts, resulting in the abortive merogony. A problem of the abortive haemosporidian development is related to health both in bird and blood-sucking insects, but remains largely unexplored [22, 27, 30, 229]. This study calls for research on the exo-erythrocytic development, which might broadly inform strategies to mitigate wildlife haemosporidioses due to identifying patterns of occurrence, pathology and mechanisms of abortion in still neglected agents of avian infection. A methodology combining the traditional histological techniques with molecular diagnostic tools is essential to speed research in this field of avian malariology. The prominent knowledge gained due to research by former generations of parasitologists provides a good starting point for addressing mechanisms of exo-erythrocytic development in haemosporidians.
Authors' contributions
GV collected published articles and collection material, analysed the literature data and wrote the manuscript; GV and TAI analysed preparations of the exo-erythrocytic stages; TAI prepared plates of images. Both authors read and approved the final manuscript.
This article benefited from comments made by Richard W. Ashford. We thank R. Adlard, E. Hoberg and A. Warren for assistance in collecting preparations of tissue stages of haemosporidian parasites, which were deposited in the International Reference Centre for Avian Haematozoa (the Queensland Museum, Queensland), in the US National Parasite Collection, Beltsville (now available in the National Museum of Natural History, Washington, DC), and Natural History Museum (London), respectively. Carter Atkinson kindly provided preparations of the megalomeronts of H. mansoni. Mikas Ilgūnas is acknowledged for technical assistance.
Competing interests
The authors declare that they have no competing interests.
Availability of data and materials
All data generated during this study are included in this published article.
This study was funded by the Research Council of Lithuania (No. MIP-045/2015).
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
Authors' Affiliations
- Santiago-Alarcon D, Palinauskas V, Schaefer HM. Diptera vectors of avian Haemosporidian parasites: untangling parasite life cycles and their taxonomy. Biol Rev Camb Philos Soc. 2012;87:928–64.PubMedView ArticleGoogle Scholar
- Garnham PCC. Malaria parasites and other Haemosporidia. Oxford: Black-well; 1966.Google Scholar
- Seed TM, Manwell RD. Plasmodia of birds. In: Kreier JP, editor. Parasitic Protozoa Vol III. Gregarines, Haemogregarines, Coccidia, Plasmodia, and Haemoproteids. New York: Academic Press; 1977. p. 311–57.Google Scholar
- Valkiūnas G. Avian malaria parasites and other Haemosporidia. Boca Raton: CRC; 2005.Google Scholar
- Atkinson CT, Thomas NJ, Hunter DB. Parasitic diseases of wild birds. Oxford: Wiley-Blackwell; 2008.View ArticleGoogle Scholar
- Telford SR. The hemoparasites of the reptilian. Boca Raton: CRC; 2009.Google Scholar
- Bray RS. Studies on the exo-erythrocytic cycle in the genus Plasmodium. Mem Lond Sch Hyg Trop Med. 1957;12:1–92.Google Scholar
- Huff CG. Exoerythrocytic stages of avian and reptilian malarial parasites. Exp Parasitol. 1969;24:383–421.PubMedView ArticleGoogle Scholar
- Marzal A. Recent advances in studies on avian malaria parasites. In: Okwa OO, editor. Malaria parasites. InTech: Croatia; 2012. p. 135–58.Google Scholar
- Frevert U, Späth GF, Yee H. Exoerythrocytic development of Plasmodium gallinaceum in the White Leghorn chicken. Int J Parasitol. 2008;38:655–72.PubMedView ArticleGoogle Scholar
- Sherman IW, editor. Malaria: parasite biology, pathogenesis, and protection. Washington: ASM; 1998.Google Scholar
- Sullivan DJ, Krishna S, editors. Malaria: drugs, disease and post-genomic biology. Berlin: Springer; 2005.Google Scholar
- Clark NJ, Clegg SM, Lima MR. A review of global diversity in avian haemosporidians (Plasmodium and Haemoproteus: Haemosporida): new insights from molecular data. Int J Parasitol. 2014;44:329–38.PubMedView ArticleGoogle Scholar
- Perkins SL. Malaria's many mates: past, present and future of the systematics of the order Haemosporida. J Parasitol. 2014;100:11–25.PubMedView ArticleGoogle Scholar
- Outlaw RK, Counterman B, Outlaw DC. Differential patterns of molecular evolution among Haemosporidian parasite groups. Parasitology. 2015;142:612–22.PubMedView ArticleGoogle Scholar
- Sehgal RN. Manifold habitat effects on the prevalence and diversity of avian blood parasites. Int J Parasitol Parasites Wildl. 2015;4:421–30.PubMedPubMed CentralView ArticleGoogle Scholar
- Bensch S, Canbäck B, DeBarry JD, Johansson T, Hellgren O, Kissinger JC, et al. The Genome of Haemoproteus tartakovskyi and its relationship to human malaria parasites. Genome Biol Evol. 2016;8:1361–73.PubMedPubMed CentralView ArticleGoogle Scholar
- Dimitrov D, Iezhova TA, Zehtindjiev P, Bobeva A, Ilieva M, Kirilova M, et al. Molecular characterisation of three avian haemoproteids (Haemosporida, Haemoproteidae), with the description of Haemoproteus (Parahaemoproteus) palloris n. sp. Syst Parasitol. 2016;93:431–49.PubMedView ArticleGoogle Scholar
- Ricklefs RE, Medeiros M, Ellis VA, Svensson-Coelho M, Blake JG, Loiselle BA, et al. Avian migration and the distribution of malaria parasites in New World passerine birds. J Biogeogr. 2016. doi:10.1111/jbi.12928.Google Scholar
- Cardona CJ, Ihejirika A, McClellan L. Haemoproteus lophortyx infection in Bobwhite Quail. Avian Dis. 2002;46:249–55.PubMedView ArticleGoogle Scholar
- Ferrell ST, Snowden K, Marlar AB, Garner M, Lung NP. Fatal hemoprotozoal infections in multiple avian species in a zoological park. J Zoo Wildl Med. 2007;38:309–16.PubMedView ArticleGoogle Scholar
- Donovan TA, Schrenzel M, Tucker TA, Pessier AP, Stalis IH. Hepatic hemorrhage, hemocoelom, and sudden death due to Haemoproteus infection in passerine birds: Eleven cases. J Vet Diagn Invest. 2008;20:304–13.PubMedView ArticleGoogle Scholar
- Olias P, Wegelin M, Zenker W, Freter S, Gruber AD, Klopfleisch R. Avian malaria deaths in parrots, Europe. Emerg Infect Dis. 2011;17:950–2.PubMedPubMed CentralView ArticleGoogle Scholar
- Pacheco MA, Escalante AA, Garner MM, Bradley GA, Aguilar RF. Haemosporidian infection in captive masked bobwhite quail (Colinus virginianus ridgwayi), an endangered subspecies of the northern bobwhite quail. Vet Parasitol. 2011;182:113–20.PubMedPubMed CentralView ArticleGoogle Scholar
- Cannell BL, Krasnec KV, Campbell K, Jones HI, Miller RD, Stephens N. The pathology and pathogenicity of a novel Haemoproteus spp. infection in wild Little Penguins (Eudyptula minor). Vet Parasitol. 2013;197:74–84.PubMedView ArticleGoogle Scholar
- Palinauskas V, Iezhova TA, Križanauskienė A, Markovets MY, Bensch S, Valkiūnas G. Molecular characterization and distribution of Haemoproteus minutus (Haemosporida, Haemoproteidae): a pathogenic avian parasite. Parasitol Int. 2013;62:358–63.PubMedView ArticleGoogle Scholar
- Valkiūnas G, Kazlauskienė R, Bernotienė R, Palinauskas V, Iezhova TA. Abortive long-lasting sporogony of two Haemoproteus species (Haemosporida, Haemoproteidae) in the mosquito Ochlerotatus cantans, with perspectives on haemosporidian vector research. Parasitol Res. 2013;112:2159–69.PubMedView ArticleGoogle Scholar
- Valkiūnas G, Kazlauskienė R, Bernotienė R, Bukauskaite D, Palinauskas V, Iezhova TA. Haemoproteus infections (Haemosporida, Haemoproteidae) kill bird-biting mosquitoes. Parasitol Res. 2014;113:1011–8.PubMedView ArticleGoogle Scholar
- Valkiūnas G, Palinauskas V, Ilgūnas M, Bukauskaite D, Dimitrov D, Bernotienė R, et al. Molecular characterization of five widespread avian haemosporidian parasites (Haemosporida), with perspectives on the PCR-based detection of haemosporidians in wildlife. Parasitol Res. 2014;113:2251–63.PubMedView ArticleGoogle Scholar
- Valkiūnas G. Haemoproteus species. In: Mehlhorn H, editor. Encyclopedia of parasitology. Berlin: Springer; 2015. p. 1–9.View ArticleGoogle Scholar
- Farmer JN. Gizzard lesions associated with Haemoproteus sacharovi infections of pigeons. Proc Iowa Acad Sci. 1964;71:537–42.Google Scholar
- Levine ND, Beamer PD, Simon J. A disease of chickens associated with Arthrocystis galli n. g, n. sp, an organism of uncertain taxonomic position. H D Srivastava Commen; 1970. p. 429–34.Google Scholar
- Borst GHA, Zwart P. An aberrant form of Leucocytozoon infection in two quaker parakeets (Myiopsitta monachus Boddaert, 1783). Z Parasitenkd. 1972;40:131–8.PubMedView ArticleGoogle Scholar
- Smith GA. Aberrant Leucocytozoon infection in parakeets. Vet Rec. 1972;91:106.PubMedView ArticleGoogle Scholar
- Walker D, Garnham PCC. Aberrant Leucocytozoon infection in parakeets. Vet Rec. 1972;91:70–2.PubMedView ArticleGoogle Scholar
- Commichau C, Jonas D. Eine durch Leukozytozoon simondi verursachte Erkrankung bei Entenküken unter besonderer Berücksichtigung des histologischen Nachweises. Zbl Vet Med B. 1977;24:662–7.View ArticleGoogle Scholar
- Opitz HM, Jakob HJ, Wiensenhuetter E, Vasandradevi V. A myopathy associated with protozoan schizonts in chickens in commercial farms in peninsular Malaysia. Avian Pathol. 1982;11:527–34.PubMedView ArticleGoogle Scholar
- Gardiner CH, Jenkins HJ, Mahoney KS. Myositis and death in bobwhites, Colinus virginianus (L.), due to hemorrhagic cysts of a haemosporozoan of undetermined taxonomic status. J Wildl Dis. 1984;20:308–18.PubMedView ArticleGoogle Scholar
- Panigrahy B, Harmon BG, Grumbles LC. Hemorrhagic disease in canaries (Serinus canarius). Avian Dis. 1984;28:536–41.PubMedView ArticleGoogle Scholar
- Gardiner CH, Fayer R, Dubey JP. Apicomplexa: Haemoproteus and Haemosporozoa of undetermined taxonomic status. In: An atlas of protozoan parasites in animal tissues. 2nd ed. Washington, DC: Armed Forces Institute of Pathology, American Registry of Pathology; 1988. p. 75–6.Google Scholar
- Earlé RA, Bastianello SS, Bennett GF, Krecek RC. Histopathology and morphology of the tissue stages of Haemoproteus columbae causing mortality in Columbiformes. Avian Pathol. 1993;22:67–80.PubMedView ArticleGoogle Scholar
- Macwhirter P. Section 7, comparative medicine and management. Chapter 43 Passeriformes. In: Ritchie BW, Harrison GJ, Harrison LR, editors. Avian medicine: principles and application. Lake Worth: Wingers Publishing; 1994. p. 1192–3.Google Scholar
- Lederer R, Adlard RD, O'Donoghue PJ. Severe pathology associated with protozoal schizonts in two pied currawongs (Strepera graculina) from Queensland. Vet Rec. 2002;150:520–2.PubMedView ArticleGoogle Scholar
- Bennett GF, Peirce MA, Ashford RW. Avian haematozoa: mortality and pathogenicity. J Nat Hist. 1993;27:993–1001.View ArticleGoogle Scholar
- Dinhopl N, Nedorost N, Mostegl MM, Weissenbacher-Lang C, Weissenböck H. In situ hybridization and sequence analysis reveal an association of Plasmodium spp. with mortalities in wild passerine birds in Austria. Parasitol Res. 2015;114:1455–62.PubMedView ArticleGoogle Scholar
- Mendes L, Pardal S, Morais J, Antunes S, Ramos JA, Peréz-Tris J, Piersma T. Hidden haemosporidian infections in Ruffs (Philomachus pugnax) staging in Northwest Europe en route from Africa to Arctic Europe. Parasitol Res. 2013;112:2037–43.PubMedView ArticleGoogle Scholar
- Dinhopl N, Mostegl MM, Richter B, Nedorost N, Maderner A, Fragner K, Weissenböck H. Application of in situ hybridization for the detection and identification of avian malaria parasites in paraffin wax-embedded tissues from captive penguins. Avian Pathol. 2011;40:315–20.PubMedPubMed CentralView ArticleGoogle Scholar
- Schoener ER, Banda M, Howe L, Castro IC, Alley MR. Avian malaria in New Zealand. NZ Vet J. 2014;62:189–98.View ArticleGoogle Scholar
- Herman CM, Greiner EC, Bennett GF, Laird M. Bibliography of the avian blood-inhabiting Protozoa. St. John's: MUN; 1976.Google Scholar
- Bennett GF, Kučera J, Woodworth-Lynas C, Whiteway M. Bibliography of the avian blood-inhabiting Protozoa. Mem Univ of Nfld Occ Pap Biol. 1981;4(Suppl 1):1–33.Google Scholar
- Bishop MA, Bennett GF. Host-parasite catalogue of the avian haematozoa: supplement 1, and Bibliography of the avian blood-inhabiting haematozoa. Mem Univ Nfld Occ Pap Biol. 1992;15(Suppl 2):1–244.Google Scholar
- Hewitt R. Bird malaria. Baltimore: Jonhs Hopkins Press; 1940.Google Scholar
- Garnham PCC. Malaria in its various vertebrate hosts. In: Kreier JP, editor. Malaria Part 1. Epidemiology, chemotherapy, morphology and metabolism. New York: Academic Press; 1980. p. 95–144.Google Scholar
- Huff CG. Observations on the preerythrocytic stages of Plasmodium relictum, Plasmodium cathemerium and Plasmodium gallinaceum in various birds. J Infect Dis. 1951;88:17–26.PubMedView ArticleGoogle Scholar
- Huff CG. Organ and tissue distribution of the exoerythrocytic stages of various avian malarial parasites. Exp Parasitol. 1957;6:143–62.PubMedView ArticleGoogle Scholar
- Raffaele G. Un ceppo italiano di Plasmodium elongatum. Riv Malar. 1934;13:3–8.Google Scholar
- James SP, Tate P. New knowledge of the life-cycle of malaria parasites. Nature. 1937;139:545.View ArticleGoogle Scholar
- James SP, Tate P. Exoerythrocytic schizogony in Plasmodium gallinaceum Brumpt, 1935. Parasitology. 1938;30:128–9.View ArticleGoogle Scholar
- Kikuth W, Mudrow L. Die endothelialen Stadien der Malariaparasiten in experiment und theorie. Zentbl Bakt Parasit K. 1938;142:113–33.Google Scholar
- Vanstreels RE, da Silva-Filho RP, Kolesnikovas CK, Bhering RC, Ruoppolo V, Epiphanio S, et al. Epidemiology and pathology of avian malaria in penguins undergoing rehabilitation in Brazil. Vet Res. 2015;46:30.PubMedPubMed CentralView ArticleGoogle Scholar
- Corradetti A. Relapses and immunological course in Plasmodium infections. Parassitologia. 1959;1:91–6.Google Scholar
- Coulston F, Cantrell W, Huff CG. The distribution and localization of sporozoites and pre-erythrocytic stages in infections with Plasmodium gallinaceum. J Infect Dis. 1945;76:226–38.View ArticleGoogle Scholar
- McGhee RB. Pre-erythrocytic development of Plasmodium gallinaceum in avian embryos. J Infect Dis. 1949;84:105–10.PubMedView ArticleGoogle Scholar
- Corradetti A, Verolini F, Neri I. Plasmodium (Haemamoeba) giovannolai n. sp. parassita di Turdus merula. Parassitologia. 1963;5:11–8.Google Scholar
- Corradetti A, Verolini F, Neri I. Studi sull'infezione da Plasmodium (Haemamoeba) giovannolai nel canarino. Parassitologia. 1963;5:73–85.Google Scholar
- Garnham PCC, Duggan AJ. Catalogue of the Garnham collection of malaria parasites and other Haemosporidia. London: The Wellcome Trust; 1986.Google Scholar
- Gabaldon A, Ulloa G. Las formas exoeritrocíticas de Plasmodium (Haemamoeba) lutzi Lucena, 1939 y presencia de esta especie en Venezuela. Bol Dir Malariol San Amb. 1976;16:299–312.Google Scholar
- Becker ER. Some unfinished investigations of malaria in pigeons. J Protozool. 1961;8:1–7.View ArticleGoogle Scholar
- Corradetti A, Neri I, Scanga M. Segnalazione in Italia di Plasmodium praecox var. matutinum in Turdus iliacus, e separazione di questo plasmodio da Plasmodium praecox come specie distinta: Plasmodium matutinum Huff, 1937. Parassitologia. 1960;2:333–43.Google Scholar
- Demina NA. Contribution à l'étude de l'influence du régime nycthéméral de l'hote sur l'évolution de la multiplication de Plasmodium relictum. Riv Malariol. 1959;38:27–44.PubMedGoogle Scholar
- Huff CG. A new variety of Plasmodium relictum from the robin. J Parasitol. 1937;23:400–4.View ArticleGoogle Scholar
- Manwell RD. Life-cycle of Plasmodium relictum var. matutinum. Am J Trop Med. 1940;20:859–66.Google Scholar
- Corradetti A, Garnham PCC, Neri L, Scanga M, Cavallini C. A redescription of Plasmodium (Haemamoeba) relictum (Grassi and Feletti, 1891). Parassitologia. 1970;12:1–10.Google Scholar
- Coulson F, Huff CG. The morphology of cryptozoites and metacryptozoites of P. relictum and the relationships of these stages to parasitaemia in canaries and pigeons. J Infect Dis. 1947;80:209–17.View ArticleGoogle Scholar
- Raffaele G. Presumibili forme iniziale di evoluzione di P. relictum. Riv Malar. 1936;15:318–24.Google Scholar
- Reichenow E, Mudrow L. Der Entwicklungsgang von P. praecox in Vogelkörper. Deutsche Tropenmedizinische Zeitschrift. 1943;46:28–299.Google Scholar
- Gabaldon A, Ulloa G. Plasmodium (Haemamoeba) tejerai sp. n. del pavo domésstico (Meleagris gallopavo) de Venezuela. Bol Dir Malariol San Amb. 1977;17:255–73.Google Scholar
- Silveira P, Belo NO, Lacorte GA, Kolesnikovas CKM, Vanstreels RET, Steindel M, Catão-Dias JL, Valkiūnas G, Braga TM. Parasitological and new molecular-phylogenetic characterization of the malaria parasite Plasmodium tejerai in South American penguins. Parasitol Int. 2013;62:165–71.PubMedView ArticleGoogle Scholar
- Vanstreels RE, Kolesnikovas CK, Sandri S, Silveira P, Belo NO, Ferreira Junior FC, et al. Outbreak of avian malaria associated to multiple species of Plasmodium in magellanic penguins undergoing rehabilitation in southern Brazil. PLoS ONE. 2014;9:e94994.PubMedPubMed CentralView ArticleGoogle Scholar
- Corradetti A, Adames A, Neri L, di Giorgi R, Cavallini C. An Italian strain of Plasmodium (Giovannolaia) circumflexum. In: Proceedings. 1st International Congress for Parasitolology. Rome; 1964. p. 238–9.Google Scholar
- Coulston F, Manwell RD. Single parasite infections and exoerythrocytic schizogony in Plasmodium circumflexum. Am J Hyg. 1941;34:119–25.Google Scholar
- Huchzermeyer FW, van der Vyver FH. Isolation of Plasmodium circumflexum from wild guineafowl (Numida meleagris) and the experimental infection in domestic poultry. Avian Pathol. 1991;20:213–23.PubMedView ArticleGoogle Scholar
- Manwell RD, Goldstein F. Life history and immunity studies of the avian malaria parasite, Plasmodium circumflexum. Proc Soc Exp Biol Med. 1938;39:426–8.View ArticleGoogle Scholar
- Paraense WL. Observaēões sōbre uma amostra brasileira do Plasmodium circumflexum. Mem Inst Oswaldo Cruz. 1952;50:211–41.PubMedView ArticleGoogle Scholar
- Huchzermeyer FW. A host-parasite list of the haematozoa of domestic poultry in sub-Saharan Africa and the isolation of Plasmodium durae Herman from turkeys and francolins in South Africa. Onderstepoort J Vet Res. 1993;60:15–21.PubMedGoogle Scholar
- Huchzermeyer FW. Pathogenicity and chemotherapy of Plasmodium durae in experimentally infected domestic turkeys. Onderstepoort J Vet Res. 1993;60:103–10.PubMedGoogle Scholar
- Laird M. Plasmodium durae Herman from the introduced common peafowl in Northern Nigeria. J Protozool. 1978;25:21–4.PubMedView ArticleGoogle Scholar
- Purchase HS. Turkey malaria. Parasitology. 1942;34:278–83.View ArticleGoogle Scholar
- Simpson ML. Exoerythrocytic stages of P. durae. J Parasitol. 1944;30:177–8.View ArticleGoogle Scholar
- Huff CG, Marchbank DF, Saroff AH, Scrimshaw PW, Shiroishi T. Experimental infections with Plasmodium fallax Schwetz isolated from the Uganda tufted guinea fowl Numida meleagris major Hartlaub. J Natl Malar Soc. 1950;9:307–19.PubMedGoogle Scholar
- Huff CG, Weathersby AB, Pipkin AC, Algire GH. The growth of exoerythrocytic stages of avian malaria within diffusion chambers in different hosts. Exp Parasitol. 1960;9:98–104.PubMedView ArticleGoogle Scholar
- Garnham PCC. A new malaria parasite of pigeons and ducks from Venezuela. Protistologica. 1977;13:113–25.Google Scholar
- Guindy E, Hoogstraal H, Mohammed AHH. Plasmodium garnhami sp. nov. from the Egyptian hoopoe (Upupa epops major Brehm). Trans R Soc Trop Med Hyg. 1965;59:280–4.PubMedView ArticleGoogle Scholar
- Ilgūnas M, Bukauskaitė D, Palinauskas V, Iezhova TA, Dinhopl N, Nedorost N, et al. Mortality and pathology in birds due to Plasmodium (Giovannolaia) homocircumflexum infection, with emphasis on the exoerythrocytic development of avian malaria parasites. Malar J. 2016;15:256.PubMedPubMed CentralView ArticleGoogle Scholar
- Huff CG, Coulston F, Laird RL, Porter RJ. Pre-erythrocytic development of Plasmodium lophurae in various hosts. J Infect Dis. 1947;81:7–13.PubMedView ArticleGoogle Scholar
- Manresa M. The occurrence of phanerozoites of Plasmodium lophurae in blood inoculated turkeys. J Parasitol. 1953;39:452–5.PubMedView ArticleGoogle Scholar
- Tonkin IM, Hawking F. Growth of Protozoa in tissue culture. IV. Plasmodium lophurae, exoerythrocytic forms, in vivo and in vitro. Trans R Soc Trop Med Hyg. 1947;41:407–17.PubMedView ArticleGoogle Scholar
- Manwell RD. Plasmodium octamerium n. sp., an avian malaria parasite from the pintail whydah bird Vidua macroura. J Protozool. 1968;15:680–5.PubMedView ArticleGoogle Scholar
- Muniz J, Soares R. Nota sōbre um parasita do genero Plasmodium encontrado no Ramphastos toco Müller, 1776, "Tugano-Aēu", e diferente do Plasmodium huffi: Plasmodium pinottii n. sp. Rev Bras Malariol. 1954;6:611–7.Google Scholar
- Ayala SC, Varela CE. Malaria in Chordeiles minor (Aves: Caprimulgidae), an intercontinental migrant. Rev Inst Med Trop Sao Paulo. 1975;17:253–6.PubMedGoogle Scholar
- Corradetti A, Scanga M. Notes on Plasmodium (Giovannolaia) polare and its transmission with Culiseta longiareolata. Parassitologia. 1965;7:61–4.Google Scholar
- Greiner EC, Black DJ, Iverson WO. Plasmodium in a bald eagle (Haliaeetus leucocephalus) in Florida. J Wildl Dis. 1981;17:555–8.PubMedView ArticleGoogle Scholar
- Telford SR, Nesbitt SA, Spalding MG, Forrester DJ. A species of Plasmodium from sandhill cranes in Florida. J Parasitol. 1994;80:497–9.PubMedView ArticleGoogle Scholar
- Gabaldon A, Ulloa G. A new species of the subgenus Novyella (Haemosporina, Plasmodiidae) from Aramides cajanea (Gruiformes, Rallidae). In: Canning EU, editor. Parasitological topics. A presentation volume to P.C.C. Garnham, F.R.S. on the occasion of his 80th birthday. 1981. p. 100–5.Google Scholar
- de Jong AC. Plasmodium dissanaikei n. sp. a new avian malaria parasite from the rose-ringed parakeet of Ceylon, Psittacula krameri manillensis. Ceylon J Med Sci. 1971;20:41–5.Google Scholar
- Manwell RD. Exoerythrocytic schizogony in Plasmodium hexamerium. Am J Hyg. 1951;53:244–8.PubMedGoogle Scholar
- Huff CG. Exoerythrocytic stages of Plasmodium nucleophilum. J Parasitol. 1956;42:612.View ArticleGoogle Scholar
- Manwell RD, Sessler GJ. Malaria parasites of toucans. J Protozool. 1971;18:570–4.PubMedView ArticleGoogle Scholar
- Manwell RD, Sessler GJ. Plasmodium paranucleophilum n. sp. from a South American tanager. J Protozool. 1971;18:629–32.PubMedView ArticleGoogle Scholar
- Corradetti A, Scanga M. Plasmodium (Novyella) vaughani subsp. merulae, n. subsp., parassita di Turdus merula, con descrizione del ciclo pre-eritrocitico. Parassitologia. 1972;14:85–93.Google Scholar
- Nelson P. Exo-erythrocytic stages of a Ceylon strain of Plasmodium vaughani in pigeons. Trans R Soc Trop Med Hyg. 1966;60:424.View ArticleGoogle Scholar
- Barretto MP. Malária Aviária: III. Sobre o encontro de formas exoeritocíticas do 'Plasmodium juxtanucreare' Versiani e Gomas, 1941. O Hospital; 1943. p. 643–5.Google Scholar
- Dhanapala SB. The occurrence of Plasmodium juxtanucleare Versiani and Gomes 1941 in domestic fowls in Ceylon. Riv Malar. 1962;41:39–46.PubMedGoogle Scholar
- Paraense WG. Observações preliminaries sobre o ciclo exoeritroocitário do Plasmodium juxtanucleare Versiani e Gomes 1941. Mems Inst Oswaldo Cruz. 1947;45:813–24.View ArticleGoogle Scholar
- Corradetti A, Neri I, Scanga M, Cavallini C. I cicli pre-eritrocitico e sporogonico di Plasmodium (Huffia) elongatum. Parassitologia. 1968;10:133–43.Google Scholar
- Garnham PCC. Patterns of exoerythrocytic schizogony. Br Med Bull. 1951;8:10–5.PubMedView ArticleGoogle Scholar
- Palinauskas V, Žiegytė R, Iezhova TA, Ilgūnas M, Bernotienė R, Valkiūnas G. Description, molecular characterisation, diagnostics and life cycle of Plasmodium elongatum (lineage pERIRUB01), the virulent avian malaria parasite. Int J Parasitol. 2016;46:697–707.PubMedView ArticleGoogle Scholar
- Raffaele G. Un ceppo italiano di Plasmodium elongatum. Riv Malariol. 1934;13:332–7.Google Scholar
- Forrester DJ, Humphrey PP. Susceptibility of the knot (Calidris canutus) to Plasmodium hermani. J Parasitol. 1981;67:747–8.PubMedView ArticleGoogle Scholar
- Forrester DJ, Nayar JK, Young MD. Natural infection of Plasmodium hermani in the northern bobwhite, Colinus virginianus, in Florida. J Parasitol. 1987;73:865–6.PubMedView ArticleGoogle Scholar
- Nayar JK, Young MD, Forrester DJ. Experimental transmission by mosquitoes of Plasmodium hermani between domestic turkeys and pen-reared bobwhites. J Parasitol. 1982;68:874–6.PubMedView ArticleGoogle Scholar
- Telford SR, Forrester DJ. Plasmodium (Huffia) hermani sp. n. from wild turkeys (Meleagris gallopavo) in Florida. J Protozool. 1975;22:324–8.View ArticleGoogle Scholar
- Huff CG. Observations on Plasmodium huffi Muniz, Soares et Batista. Am J Trop Med Hyg. 1953;2:620–3.PubMedGoogle Scholar
- Muniz J, Soares R, Batista S. Sōbre uma espécie de Plasmodium parasita do Ramphastos toco Müller, 1776. Plasmodium huffi n. sp. Rev Bras Malariol. 1951;3:339–44.Google Scholar
- Grilo ML, Vanstreels RET, Wallace R, García-Párraga D, Braga EM, Chitty J, et al. Malaria in penguins–current perceptions. Avian Pathol. 2016;45:393–407.PubMedView ArticleGoogle Scholar
- Howe L, Castro IC, Schoener ER, Hunter S, Barraclough RK, Alley MR. Malaria parasites (Plasmodium spp.) infecting introduced, native and endemic New Zealand birds. Parasitol Res. 2012;110:913–23.PubMedView ArticleGoogle Scholar
- Palinauskas V, Žiegytė R, Ilgūnas M, Iezhova TA, Bernotienė R, Bolshakov C, et al. Description of the first cryptic avian malaria parasite, Plasmodium homocircumflexum n. sp. with experimental data on its virulence and development in avian hosts and mosquitoes. Int J Parasitol. 2015;45:51–62.PubMedView ArticleGoogle Scholar
- Permin A, Juhl J. The development of Plasmodium gallinaceum infections in chickens following single infections with three different dose levels. Vet Parasitol. 2002;105:1–10.PubMedView ArticleGoogle Scholar
- Fleischman RW, Squire RA, Sladen WJL, Melby EC. Malaria (Plasmodium elongatum) in captive African penguins (Spheniscus demersus). J Am Vet Med Assoc. 1968;153:928–35.PubMedGoogle Scholar
- Porter RJ. The tissue distribution of exoerythrocytic schizonts in sporozoites-induces infections with Plasmodium cathemerium. J Infect Dis. 1942;71:1–17.View ArticleGoogle Scholar
- Hegner R, Wolfson F. Association of Plasmodium and Toxoplasma-like parasites in birds. Am J Hyg. 1938;28:437–54.Google Scholar
- Mohammed AHH. Systematic and experimental studies on protozoan blood parasites of Egyptian birds. Cairo: Cairo University Press; 1958.Google Scholar
- Valkiūnas G, Ilgūnas M, Bukauskaitė D, Žiegytė R, Bernotienė R, Jusys V, et al. Plasmodium delichoni n. sp.: description, molecular characterisation and remarks on the exoerythrocytic merogony, persistence, vectors and transmission. Parasitol Res. 2016;115:2625–36.PubMedView ArticleGoogle Scholar
- Valkiūnas G, Iezhova TA, Loiseau C, Smith TB, Sehgal RNM. New malaria parasites of the subgenus Novyella in African rainforest birds, with remarks on their high prevalence, classification and diagnostics. Parasitol Res. 2009;104:1061–77.PubMedView ArticleGoogle Scholar
- Mantilla JS, González AD, Valkiūnas G, Moncada LI, Matta NE. Description and molecular characterization of Plasmodium (Novyella) unalis sp. nov. from the Great Thrush (Turdus fuscater) in highland of Colombia. Parasitol Res. 2013;112:4193–204.PubMedView ArticleGoogle Scholar
- Holding PA, Snow RW. Impact of Plasmodium falciparum malaria on performance and learning: review of the evidence. Am J Trop Med Hyg. 2001;64(Suppl 1–2):68–75.PubMedGoogle Scholar
- Adams Y, Kuhnrae P, Higgins MK, Ghumra A, Rowe JA. Rosetting Plasmodium falciparum-infected erythrocytes bind to human brain microvascular endothelial cells in vitro, demonstrating a dual adhesion phenotype mediated by distinct P. falciparum erythrocyte membrane protein 1 domains. Infect Immun. 2014;82:949–59.PubMedPubMed CentralView ArticleGoogle Scholar
- Lainson R, Landau I, Shaw JJ. On a new family of non-pigmented parasites in the blood of reptiles: Garniidae fam. nov., (Coccidiida: Haemosporidiidea). Some species of the new genus Garnia. Int J Parasitol. 1971;1:241–50.View ArticleGoogle Scholar
- Lainson R. Atlas of protozoan parasites of the Amazonian fauna of Brazil. Vol. 1. Haemosporida of reptiles, vol. 1. Ananindeua: Instituto Evandro Chagas; 2012.Google Scholar
- Amoudi MA, Alyousif MS, Saifi MA, Alanazi AD. A new species of plasmodiidae (Coccidia: Hemosporidia) from the blood of the skink Scincus hemprichii (Scincidae: Reptilia) in Saudi Arabia. Saudi J Biol Sci. 2015;22:312–6.PubMedView ArticleGoogle Scholar
- Perkins SL. Species concepts and malaria parasites: detecting a cryptic species of Plasmodium. Proc Biol Sci. 2000;267:2345–50.PubMedPubMed CentralView ArticleGoogle Scholar
- Gabaldon A, Ulloa G, Zerpa N. Fallisia (Plasmodioides) neotropicalis subgen. nov. sp. nov. from Venezuela. Parasitology. 1985;90:217–25.View ArticleGoogle Scholar
- Outlaw DC, Ricklefs RE. Species limits in avian malaria parasites (Haemosporida): how to move forward in the molecular era. Parasitology. 2014;141:1223–32.PubMedView ArticleGoogle Scholar
- Levin II, Valkiūnas G, Iezhova TA, O'Brien SL, Parker PG. Novel Haemoproteus species (Haemosporida: Haemoproteidae) from the swallow-tailed gull (Lariidae), with remarks on the host range of hippoboscid-transmitted avian hemoproteids. J Parasitol. 2012;98:847–54.PubMedView ArticleGoogle Scholar
- Iezhova T. Comparative morphology of haemoproteids (Haemosporidia, Haemoproteidae) of the Palearctic birds (in Russian). Ph.D. Thesis. Vilnius: Institute of Ecology; 1994.Google Scholar
- Peirce MA. Haemoproteus balearicae sp. nov., from crowned cranes, Balearica pavonina pavonina and B. pavonina gibbericeps. Bull Epizoot Dis Afr. 1973;21:467–75.PubMedGoogle Scholar
- Khan RA, Fallis AM. Endogenous stages of Parahaemoproteus fringillae (Labbé, 1894) and Leucocytozoon fringillinarum Woodcock, 1910. Can J Zool. 1969;47:37–9.View ArticleGoogle Scholar
- Peirce MA, Lederer R, Adlard RD, O'Donoghue PJ. Pathology associated with endogenous development of haematozoa in birds from southeast Queensland. Avian Pathol. 2004;33:445–50.PubMedView ArticleGoogle Scholar
- Miltgen F, Landau I, Ratanaworabhan N, Yenbutra S. Parahaemoproteus desseri n. sp.; Gamétogonie et schizogonie chez l'hôte naturel: Psittacula roseata de Thailande, et sporogonie expérimentale chez Culicoides nubeculosus. Ann Parasitol Hum Comp. 1981;56:123–30.PubMedGoogle Scholar
- Atkinson CT, Greiner EC, Forrester DJ. Pre-erythrocytic development and associated host responses to Haemoproteus meleagridis (Haemosporina: Haemoproteidae) in experimentally infected domestic turkeys. J Protozool. 1986;33:375–81.PubMedView ArticleGoogle Scholar
- Atkinson CT, Forrester DJ, Greiner EC. Pathogenicity of Haemoproteus meleagridis (Haemosporina: Haemoproteidae) in experimentally infected domestic turkeys. J Protozool. 1988;74:228–39.Google Scholar
- Sibley LD, Werner JK. Susceptibility of pekin and muscovy ducks to Haemoproteus nettionis. J Wildl Dis. 1984;20:108–13.PubMedView ArticleGoogle Scholar
- Anschütz G. Ueber den Entwickelungsgang des "Haemoproteus orizivorae" nov. spec. Zentralbl Bakteriol Parasitenkd Infektionskr Hyg I Abt Orig. 1909;51:654–9.Google Scholar
- Burtikashvili LP. New species of blood parasite, Haemoproteus zasukhini sp. nov. (Sporozoa, Haemoproteidae) from the house sparrow (Passer domesticus L.) (in Russian). Soobsh Akad Nauk Gruz SSR. 1973;72:697–700.Google Scholar
- Burtikashvili LP. Blood parasites of wild birds in Georgia. Tbilisi: Metsniereba; 1978 (In Russian).Google Scholar
- Peirce MA. Haemoproteid parasites of Passer spp. Parasitology. 1976;73:407–15.PubMedView ArticleGoogle Scholar
- Wenyon CM. Protozoology: a manual for medical men, veterinarians and zoologists, etc. London: Bailliére Tindal & Cox; 1926.View ArticleGoogle Scholar
- Garvin MC, Homer BL, Greiner EC. Pathogenicity of Haemoproteus danilewskyi, Kruse, 1890, in Blue Jays (Cyanocitta cristata). J Wildl Dis. 2003;39:161–9.PubMedView ArticleGoogle Scholar
- Ahmed FE, Mohammed AHH. Schizogony in Haemoproteus columbae Kruse. J Protozool. 1977;24:389–93.PubMedView ArticleGoogle Scholar
- Acton HW, Knowles R. Studies on the halteridium parasite of the pigeon, Haemoproteus columbae Celli and San Felice. Indian J Med Res. 1914;1:663–90.Google Scholar
- Aragão HB. Über den Entwicklungsgang and die Übertragung von Haemoproteus columbae. Arch Protistenkd. 1908;12:154–67.Google Scholar
- Huff CG. Schizogony and gametocyte development in Leucocytozoon simondi, and comparison with Plasmodium and Haemoproteus. J Infect Dis. 1942;71:18–32.View ArticleGoogle Scholar
- Linxian H, Yuguang L, Hongping J. On sporogony and schizogony of Haemoproteus columbae of pigeon. Acta Zool Sinica. 1989;35:306–12.Google Scholar
- Mohammed AHH. Studies on the schizogony of Haemoproteus columbae Kruse, 1890. Proc Egypt Acad Sci. 1967;19:37–46.Google Scholar
- Baker JR. Haemoproteus palumbis sp. nov. (Sporozoa, Haemosporina) of the English Wood-Pigeon Columba p. palumbus. J Protozool. 1966;13:515–9.PubMedView ArticleGoogle Scholar
- Desser SS, Bennett GF. The genera Leucocytozoon, Haemoproteus and Hepatocystis. In: Kreier JP, Baker JR, editors. Parasitic protozoa. 2nd ed. Cambridge: Academic Press; 1993. p. 273–307.View ArticleGoogle Scholar
- Mutlow A, Forbes N. Haemoproteus in raptors: pathogenicity, treatment and control. Wallbridge: Landsdown Veterinary Surgeons; 1999.Google Scholar
- Paperna I, Gil H. Schizogonic stages of Haemoproteus from Wenyon's Baghdad sparrows are also found in Passer domesticus biblicus in Israel. Parasitol Res. 2003;91:486–90.PubMedView ArticleGoogle Scholar
- Fowler NG, Forbes GB. Aberrant Leucocytozoon infection in parakeets. Vet Rec. 1972;91:345–7.PubMedView ArticleGoogle Scholar
- Markus MB. Leucocytozoon infection in parakeets. Vet Res. 1972;91:323.Google Scholar
- Garnham PCC. Epizootics of Leucocytozoon infections in parakeets in England. In: Progress in parasitology: abstracts of papers read at the 4-th International Congress; 1973.Google Scholar
- Simpson VR. Leucocytozoon-like infections in parakeets, budgeries and a common buzzard. Vet Rec. 1991;13:30–2.View ArticleGoogle Scholar
- Forrester DJ, Greiner EC. Leucocytozoonosis. In: Atkinson CT, Thomas NJ, Hunter DB, editors. Parasitic diseases of wild birds. Oxford: Wiley-Blackwell; 2008. p. 54–107.Google Scholar
- Peirce MA, Adlard RD, Lederer R. A new species of Leucocytozoon Berestneff, 1904 (Apicomplexa: Leucocytozoidae) from the avian family Artamidae. Syst Parasitol. 2005;60:151–4.PubMedView ArticleGoogle Scholar
- Clark GW. Schizogony and gametocyte development of Leucocytozoon berestneffi in the yellow-billed magpie Pica nuttalli. J Protozool. 1965;12:584–9.View ArticleGoogle Scholar
- Khan RA, Fallis AM. Speciation, transmission, and schizogony of Leucocytozoon in corvid birds. Can J Zool. 1971;49:1363–7.PubMedView ArticleGoogle Scholar
- Khan RA. Development of Leucocytozoon ziemanni (Laveran). J Parasitol. 1975;61:449–57.View ArticleGoogle Scholar
- Wong TC, Desser SS. An ultrastructural study of the schizogonic development of Leucocytozoon dubreuili in the liver of the American robin, Turdus migratorius. Protistologica. 1981;17:11–27.Google Scholar
- Khan RA, Fallis AM. Life cycles of Leucocytozoon dubreuili Mathis and Léger, 1911 and L. fringillinarum Woodcock, 1910 (Haemosporidia: Leucocytozoidae). J Protozool. 1970;17:642–58.View ArticleGoogle Scholar
- Wong TC, Desser SS. Ultrastructural observations on renal schizogony of Leucocytozoon dubreuili in the American robin. J Protozool. 1978;25:302–14.PubMedView ArticleGoogle Scholar
- Borg K. On Leucocytozoon is Swedish capercaillie, black grouse and hazel grouse. Lund: Lund Publisher; 1953.Google Scholar
- Clarke CHD. Organisms of a malarial type in ruffed grouse, with a description of the schizogony of Leucocytozoon bonasae. J Wildl Manag. 1938;2:146–50.View ArticleGoogle Scholar
- Newman MW. Large schizonts of Leucocytozoon from the kidney of the ruffed grouse (Bonasa umbellus). Can J Zool. 1970;48:879–80.View ArticleGoogle Scholar
- Hong L, Chen X, Lin Y. The sporogony and schizogony of Leucocytozoon sabrazesi from chicken in China. Bull Soc Fr Parasitol. 1990;8(Suppl. 1):321.Google Scholar
- de Jong AC. Leucocytozoon neavei in francolins, Francolinus (=Pternistis) leucoscepus. Ceylon Vet J. 1971;19:92–5.Google Scholar
- Peirce MA, Cheke AS, Cheke RA. A survey of blood parasites of birds in the Mascarene Islands, Indian Ocean with descriptions of two new species and taxonomic discussion. Ibis. 1977;119:451–61.View ArticleGoogle Scholar
- Adlard RD, Peirce MA, Lederer R. New species of Leucocytozoon from the avian families Otidae, Podargidae and Threskiornithidae. J Nat Hist. 2002;36:1261–7.View ArticleGoogle Scholar
- Baker JR. Transmission of Leucocytozoon sakharoffi in England by Simulium angustitarse. Parasitology. 1970;60:417–23.View ArticleGoogle Scholar
- Baker JR. Sporogony and schizogony of Leucocytozoon sakharoffi in England. J Protozool. 1971;18(Suppl):40.Google Scholar
- Baker JR. Protozoan parasites of the blood of British wild birds and mammals. J Zool. 1974;172:169–90.View ArticleGoogle Scholar
- Wingstrand KG. On some haematozoa of Swedish birds with remarks on the schizogony of Leucocytozoon sakharoffi. Kungl Sven Vetenskapsakad Handl. 1947;24:1–31.Google Scholar
- Wingstrand KG. Further studies on Leucocytozoon sakharoffi. Kungl Sven Vetenskapsakad Handl. 1948;24:1–17.Google Scholar
- Chernin E. The relapse phenomenon in the Leucocytozoon simondi infection of the domestic duck. Am J Hyg. 1952;56:101–18.PubMedGoogle Scholar
- Cowan AB. Reactions against the megaloschizonts of Leucocytozoon simondi Mathis and Léger in ducks. J Infect Dis. 1957;100:82–7.PubMedView ArticleGoogle Scholar
- Desser SS. Schizogony and gametogony of Leucocytozoon simondi and associated reactions in the avian host. J Protozool. 1967;14:244–54.PubMedView ArticleGoogle Scholar
- Desser SS. The fine structure of Leucocytozoon simondi II. Megaloschizogony. Can J Zool. 1970;48:417–21.PubMedView ArticleGoogle Scholar
- Desser SS. The fine structure of Leucocytozoon simondi. VI. Hepatic schizogony. Can J Zool. 1973;51:605–9.PubMedView ArticleGoogle Scholar
- Desser SS, Fallis AM. The cytological development and encapsulation of megaloschizonts of Leucocytozoon simondi. Can J Zool. 1967;45:1061–5.PubMedView ArticleGoogle Scholar
- Desser SS, Fallis AM, Garnham PCC. Relapses in ducks chronically infected with Leucocytozoon simondi and Parahaemoproteus nettionis. Can J Zool. 1968;46:281–5.PubMedView ArticleGoogle Scholar
- Desser SS, Stuht J, Fallis AM. Leucocytozoonosis in Canada geese in Upper Michigan. I. Strain differences among geese from different localities. J Wildl Dis. 1978;14:124–31.PubMedView ArticleGoogle Scholar
- Desser SS, Ryckman AK. The development and pathogenesis of Leucocytozoon simondi in Canada and domestic geese in Algonquin Park, Ontario. Can J Zool. 1976;54:634–43.PubMedView ArticleGoogle Scholar
- Eide A, Fallis AM. Experimental studies of the life cycle of Leucocytozoon simondi in ducks in Norway. J Protozool. 1972;19:414–6.PubMedView ArticleGoogle Scholar
- Fallis AM, Davies DM, Vickers MA. Life history of Leucocytozoon simondi Mathis and Léger in natural and experimental infections and blood changes produced in the avian host. Can J Zool. 1951;29:305–28.View ArticleGoogle Scholar
- Newberne JW. Studies on the histopathology of Leucocytozoon simondi infection. Am J Vet Res. 1957;18:191–9.PubMedGoogle Scholar
- Karstad L. A case of leucocytozoonosis in a wild mallard. Bull Wildl Dis Ass. 1965;1:33–4.Google Scholar
- Khan RA, Desser SS, Fallis AM. Survival of sporozoites of Leucocytozoon in birds for 11 days. Can J Zool. 1969;47:347–50.View ArticleGoogle Scholar
- O'Roke EC. A malaria-like disease of ducks caused by Leucocytozoon anatis Wickware. Univ Mich Sch Cons Bull. 1934;4:1–44.Google Scholar
- Yang YJ. In vitro cultivation of exoerythrocytic stages of Leucocytozoon simondi. Can J Zool. 1971;49:561–3.PubMedView ArticleGoogle Scholar
- Yang YJ, Desser SS, Fallis AM. Elongate and round gametocytes of Leucocytozoon simondi (Mathis et Léger) in ducks inoculated with megaloschizonts. J Protozool. 1971;18:553–6.PubMedView ArticleGoogle Scholar
- Newberne JW. The pathology of Leucocytozoon infection in turkeys with a note on its tissue stages. Am J Vet Res. 1955;16:593–7.PubMedGoogle Scholar
- Steele EJ, Noblet GP. Schizogonic development of Leucocytozoon smithi. J Protozool. 1992;39:530–6.PubMedView ArticleGoogle Scholar
- Richey DJ, Ware RE. Schizonts of Leucocytozoon smithi in artificially infected turkeys. Cornell Vet. 1955;45:642–3.PubMedGoogle Scholar
- Wehr EE. Studies on leucocytozoonosis of turkeys, with notes on schizogony, transmission, and control of Leucocytozoon smithi. Avian Dis. 1962;6:95–210.View ArticleGoogle Scholar
- Allison FR, Desser SS. Observations on the life cycle of the haemosporidian Leucocytozoon tawaki in the simuliid vector and the New Zealand fiordland crested penguin, Eudyptes pachyrhynchus. In: Short commun. 4th International Congress of Parasitology. Warsaw; 1978.Google Scholar
- Fallis AM, Bisset SA, Allison FR. Leucocytozoon tawaki n. sp. (Eucoccida: Leucocytozoidae) from the penguin Eudyptes pachyrhynchus, and preliminary observations on its development in Austrosimulium spp. (Diptera: Simuliidae). NZ J Zool. 1976;3:11–6.View ArticleGoogle Scholar
- Akiba K. Leucocytozoonosis of chickens. Nat Inst Anim Hlth Q. 1970;10(Suppl):131–47.Google Scholar
- Akiba K, Kawashima H, Inui S, Ishii S. Studies on Leucocytozoon of chickens in Japan. I. Natural infection of L. caulleryi. Bull Natl Inst Anim Hlth. 1958;34:163–80.Google Scholar
- Akiba K, Inui S, Ishitani R. Morphology and distribution of intracellular schizonts in chickens experimentally infected with Akiba caulleryi. Natl Inst Anim Hlth Q. 1971;11:109–21.Google Scholar
- Chew M. Megaloschizonts of Leucocytozoon in the eyes and sciatic nerves of the domestic fowl. Vet Rec. 1968;83:518–9.PubMedView ArticleGoogle Scholar
- Kitaoka S, Morii T, Fujisaki K. Quantitative observation on schizogony of Akiba caulleryi in chickens. Nat Inst Anim Hlth Q. 1972;12:151–60.Google Scholar
- Lee HR, Koo BS, Jeon EO, Han MS, Min KCh, Lee SB, Bae Y, Mo IP. Pathology and molecular characterization of recent Leucocytozoon caulleryi cases in layer flocks. J Biomed Res. 2016;30:517–24.PubMedPubMed CentralView ArticleGoogle Scholar
- Omar AR. Haemoprotozoan infections of poultry in Malaysia. Kajian Vet. 1968;1:109–24.Google Scholar
- Morii T. A review of Leucocytozoon caulleryi infection in chickens. J Protozool Res. 1992;2:128–33.Google Scholar
- Morii T, Nakamura K, Lee YC, Iijima T, Hoji K. Observations on the Taiwanese strain of Leucocytozoon caulleryi (Haemosporina) in chickens. J Protozool. 1986;33:231–4.PubMedView ArticleGoogle Scholar
- Morii T, Matsui T, Kobayashi F, Iijima T, Fukuda M. Freeze fracture study of the schizonts and merozoites of Leucocytozoon caulleryi. Zbl Bakt Hyg A. 1987;264:527–35.Google Scholar
- Morii T, Fukuda M. Observations on first-generation schizogony of Leucocytozoon caulleryi in chickens. J Protozool. 1992;39:281–7.View ArticleGoogle Scholar
- Takamatsu H, Fujisaki K, Kitaoka S. Development of Leucocytozoon caulleryi in chick embryos infected by biting of Culicoides arakawae through shell membrane. Natl Inst Anim Health Q. 1978;18:63–8.Google Scholar
- Desser SS, Baker JR, Lake P. The fine structure of Leucocytozoon simondi. I. Gametogenesis. Can J Zool. 1970;48:331–6.PubMedView ArticleGoogle Scholar
- Bukauskaitė D, Bernotienė R, Iezhova TA, Valkiūnas G. Mechanisms of mortality in Culicoides biting midges due to Haemoproteus infection. Parasitology. 2016;143:1748–54.PubMedView ArticleGoogle Scholar
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text:
http://www.leadership-principles.com/en/ and about his book at http://amzn.to/1fW2lmX
He's worked in IT for over 20 years and is a specialist in Transformational Leadership, Operational Performance Improvement, Organisational Development, Creating Business Value via IT, and Program and Change Management.
Deb Bailey: Thanks for stopping by the blog today, Gordon. What prompted you to write your book?
Gordon Tredgold: Glad to be here, Deb. I've always wanted to write a book on leadership I think I have a very simple approach to leadership, which people appreciate and I wanted to share it. Too often I find books on leadership are overly complex and I wanted to write something simple and accessible, that could be easily understood by both leaders and aspiring leader.
I had been writing a Leadership blog, as I wasn't sure I could write a full book, then a friend who was going through chemo told me that it wasn't working and I wanted to do something to help so I decided to run a marathon. As I started to train for the marathon I realized much of what I was talking about on the blog I was applying to myself to get me to run the marathon, so I thought why not write a book combining the 2 and look to raise money for charity at the same time.
Deb: Please share what your book is about.
Gordon: It's about my approach to leadership, the principles that I look to try and apply in a business settings, and also how I applied it to myself in order to get myself as an unfit, 53 year old non-runner to run a marathon. I'm like my own guinea pig for my approach to leadership.
It's not really a book about running, its more about the journey.
Deb: That's terrific. So, what is the key message you want to deliver to your readers?
Gordon: The key message is that good leadership can deliver sustainable success, but it will take time, its not a quick fix. Success gained too easily is lost just as easily. You need to undergo a culture change to build a winning culture within an organization, and then there is no limit to the success that can be achieved,
Deb: In your book you mention "the happy underachiever," could you share more about that?
Gordon: Too often departments are content with their performance, sometimes they know they are underperforming but don't care because they are not inspired by their leader, or it could be that they don't believe they could do better, and consequently they are happy where they are, either conscious or unconscious under achiever.
Usually it requires a new leader to come in and take over, especially if the leader has been in place while as they are now part of the problem.
We can all do much more and better than we believe and as leaders we need to show people their true potential and then inspire them to look to achieve it.
I met with someone just recently who had run 66 Ultra Marathons, for charity, on consecutive days. I was very impressed I thought that was amazing, but he told me he did Ultras (50km) run because a guy from Belgian Stefan Engels was looking to run 365 marathons on consecutive days.
I have just run 2 marathons 6 months apart so I thought that was a lot, but then you hear of a guy who has run 66 Ultras, and then someone who has run 365, now I know I can do so much more.
Shortly after hearing this I read about a Spanish guy who had just run 607 marathons on consecutive days.
So even though I thought I was a happy underachiever before my marathon and became happy achiever on completing it, now I again feel like an happy under achiever as their achievements really do dwarf my own, and I now know that I am capable of so much more.
And this is true of all of us, in whatever it is we are undertaking be it business or private life.
Deb: I totally agree! Can you elaborate on what goes into creating a "culture of success?"
Gordon: The key ingredients to creating a culture of success are small successes and reward and recognition. You need to create a path of success starting small and moving onto bigger and bigger successes, slowly raising the bar and at the same time providing reward and recognition for your teams. Get them used to the feeling of being successful and then slowly raise the bar, and as the team gets into the winning habit they themselves will want to look to improve and enjoy further successes.
Its just like teaching a child to swim, you encourage them even when the can't swim, ad you just keep stepping further and further back, all the while encouraging them and helping them to feel safe, and then eventually they are swimming.
But too often, leaders and managers who know how to do this with their children, don't take the same approach with their teams. They expect them to be able to jump in at the deep end and be able to swim right off the bat. Won't happen often!
Deb: What's the best advice you could give to women entrepreneurs specifically?
Gordon: Be bold, go after big targets, everyone loves to achieve big goals, this is what inspires people. Show them the way to be successful and then encourage people along that journey. Emotional intelligence is key, and in my experience woman are better at it than men, but often they tend to play it down, I don't know why but I would play it up.
Deb: Good point. It's certainly better to lead with our strengths. You're sharing a lot of very important information in your book. What inspires you to do the work you do?
Gordon: Before I was often told that I wouldn't be successful at this, or that project was too tough, and what inspired me then was the look on peoples faces when I achieved something that they thought was impossible.
What inspires me know is the looks on the faces of my teams when they have achieved something they didn't feel was possible.
So in the marathon I was happy to complete it, and prove people who told me I couldn't do it they were wrong. But what pleased me much more was the look on Taraks face when he completed Dusseldorf Marathon with me, and also the looks on my sister Alyson and my colleagues Dave and Bart when they completed the Cologne Marathon. They we're marathons runners, they didn't think they could do it, but the triumphed, and the look on their faces at the end was worth all of the effort involved.
Deb: So glad you could share your book with us. Where can readers find it?
Gordon: Thanks for featuring it. You can find my book on Amazon.com
Print - http://www.amazon.com/Leadership-Marathon-Everything-Sustainable-Achievements/dp/190962327X/
Kindle - http://www.amazon.com/Leadership-Its-Marathon-Not-Sprint-ebook/dp/B00H2ZE2SM/
Or if you want a signed copy you can find it on: www.leadership-its-a-marathon-not-a-sprint.com
Deb: Please share your website and social media URLs.
Gordon: Here you go...
My blog: www.leadership-principles.com
Facebook: www.facebook.com/leadershipprinciples
Twitter: @gordontredgold
Author Q&A: Leadership: It's a Marathon Not a Sprint by Gordon Tredgold
Posted by Deborah A Bailey
2 comments
- Gordon Tredgold said...
Deb, thanks for hosting me it was a great pleasure talking with you, i like to write about leadership, but i love to to about it and answer questions, it really is a passion of mine.
thanks also for providing the links to my blog and book, its really appreciated\\Gordon
- February 6, 2014 at 5:56 PM
- Deb Bailey said...
You're welcome, Gordon! I'm sure your book will resonate with my readers. Thanks for sharing your book with us.
- February 6, 2014 at 6:33 PM
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