Background: Malaria parasites (genus Plasmodium) are widespread in birds. These pathogens cause pathology of blood and various organs, often resulting in severe avian malaria. Numerous recent studies have reported DNA sequences of avian malaria parasites, indicating rich genetic diversity and the possible existence of many undescribed species. However, the majority of reported Plasmodium lineages remain unidentified to species level, and molecular characterization is unavailable for the majority of described Plasmodium parasites. During the past 15 years, numerous new Plasmodium species have been described. However, keys for their identification are unavailable or incomplete. Identification of avian malaria parasites remains a difficult task even for experts, and this precludes development of avian malariology, particularly in wildlife. Here, keys for avian malaria parasites have been developed as a baseline for assisting academic and veterinary medicine researchers in identification of these pathogens. The main obstacles and future research priorities have been defined in the taxonomy of avian Plasmodium species. Methods: The data were considered from published articles and type and voucher material, which was accessed in museums in Europe, the USA and Australia. Blood films containing various blood stages of the majority of described species were examined and used for the development of dichotomous keys for avian Plasmodium species. Results: In all, 164 published articles were included in this review. Blood stages of avian Plasmodium parasites belonging to subgenera Haemamoeba, Giovannolaia, Novyella, Bennettinia and Huffia were analysed and compared. Illustrated keys for identification of subgenera and species of these parasites were developed. Lists of invalid and synonymous Plasmodium parasite names as well as names of doubtful identity were composed. Conclusion: This study shows that 55 described species of avian Plasmodium can be readily identified using mor - phological features of their blood stages. These were incorporated in the keys. Numerous synonymous names of Plasmodium species and also the names belonging to the category species inquirenda exist, and they can be used as reserves for future taxonomy studies. Molecular markers are unavailable for 58% of described Plasmodium parasites, raising a task for the current avian malaria researchers to fill up this gap. Keywords: Avian malaria, Key to species, Plasmodium, Species inquirenda, Synonym, Avian Plasmodium taxonomy Background genera (Culex, Coquillettidia, Aedes, Mansonia, Culi- Malaria parasites of the genus Plasmodium (Haemospor- setta, Anopheles, Psorophora) for completing sporogony ida, Plasmodiidae) inhabit all major groups of terrestrial and transmission [1, 8–11]. This is not the case in mam - vertebrates. Avian malaria parasites is a peculiar group malian malaria parasites whose are transmitted mostly by among them, particularly due to the ability of numerous Anopheles species [1, 12–14]. Furthermore, sporogony species to develop and complete life cycles in numer- of many avian Plasmodium parasites is completed rela- ous bird species belonging to different families and even tively fast in susceptible vectors at relatively low temper- orders [1–7]. The same is true for invertebrate hosts (vec - atures [1, 8, 15, 16]. These features likely contributed to tors) of these parasites [8, 9]. Many species of avian Plas- the global distribution of some avian malaria infections, modium use Culicidae mosquitoes belonging to different which are actively transmitted in countries with warm and cold climates, including regions close to the Polar Circles [6, 17–19]. *Correspondence: email@example.com Nature Research Centre, Akademijos 2, 08412 Vilnius 2100, Lithuania © The Author(s) 2018. This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creat iveco mmons .org/licen ses/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://creat iveco mmons .org/ publi cdoma in/zero/1.0/) applies to the data made available in this article, unless otherwise stated. Valkiūnas and Iezhova Malar J (2018) 17:212 Page 2 of 24 Life cycles of avian malaria parasites are similar in their parasites [1, 12, 42]. This raises questions about para - basic features to those of human and other mammal site species identification if the same pathogen is found Plasmodium species [1, 2, 8, 13, 14, 20]. Malaria para- in unusual avian hosts. Molecular characterization is sites are obligate heteroxenous protists, with merogony helpful in diagnosis of malaria infections, and has been in cells of fixed tissues and also blood cells. Gametogony developed for detection of some avian Plasmodium spe- occurs in red blood cells, and sexual process and sporo- cies [21, 40]. Molecular markers are essential in diagnosis gony are completed in Culicidae mosquitoes. However, and identification of exo-erythrocytic and vector stages, the life cycles of avian Plasmodium species differ from which cannot be identified using morphological features those of the parasites of mammals, particularly due to [11, 43, 44]. However, molecular diagnostics using gen- their relatively low host specificity and marked varia - eral primers (the main diagnostic tool currently used in tion in patterns of development in avian hosts and vec- wildlife malariology) is often insensitive in distinguishing tors. For example, Plasmodium (Haemamoeba) relictum of avian Plasmodium spp. co-infections, which are com- infects and completes its life cycle in birds belonging to mon and even predominate in many bird populations over 300 species and 11 orders, and Plasmodium (Huffia) [45–48]. Specific molecular markers for the majority of elongatum, Plasmodium (Novyella) vaughani and many avian Plasmodium species have not been developed, and other species also have a broad range of avian hosts [6, currently are difficult to develop due to significant genetic 8, 21–23]. Erythrocytic merozoites of many avian malaria diversity of malaria parasites, which remain undescribed parasites can induce secondary tissue merogony in birds in wildlife. Morphological identification using micro - [24, 25]. The exo-erythrocytic merogony occurs in cells scopic examination of blood films remains important in of the reticuloendothelial and haemopoietic systems, malaria diagnostics in the wild, and is particularly valu- but has not been reported in hepatocytes [2, 4, 8, 23, 26]. able if it is applied in parallel with polymerase chain reac- Pedunculated oocysts were discovered in Plasmodium tion (PCR)-based diagnostic tools [5, 30, 49, 50]. (Bennettinia) juxtanucleare; these oocysts possess leg- During the past 15 years, numerous avian Plasmodium like outgrowths which attach the oocysts to the mosquito parasites were named and described using morphologi- midgut wall . These and some other features are not cal features of their blood stages [49, 51–59]. However, characteristics of malaria parasites of mammals, and this molecular markers for parasite detection were developed is reflected in genetic differences between these groups of in a handful of these descriptions. The keys that are avail - parasites and their different position in molecular phy - able for identification of avian Plasmodium species , logenies [28–33]. should be reworked in the light of the newly available Malaria, the disease caused by parasites of the genus information. Plasmodium, has traditionally been viewed as a disease of The main aim of this review is to develop easy-to-use the blood and blood forming tissues of vertebrate hosts, keys for identification of avian malaria parasites using with exo-erythrocytic stages of development causing lit- morphological features of their blood stages as a baseline tle or no pathology [1, 13, 14, 34]. While available evi- for assisting academic and veterinary medicine research- dence still supports this view for the primate and rodent ers in identification of these pathogens. Lists of synony - malarial parasites, there is increasing evidence that the mous names of Plasmodium species as well as invalid pathogenicity of tissue stages of avian species of Plasmo- species names were updated and compiled. The Plas - dium has been significantly underestimated . Even modium parasite names of unknown taxonomic posi- more, avian malaria is often a more severe disease than tion (incertae sedis) and the species of doubtful identity human malaria. There is recent experimental evidence of requiring further investigation (species inquirenda) were unexpected pathology associated with obstructive devel- specified as well. The information about useful molecular opment of secondary exo-erythrocytic stages of Plasmo- markers, which can be used for described Plasmodium dium in brain capillaries that can lead to ischaemia and species detection and comparison was also summarized. rapid death in birds that have very low intensity parasi- This review might be helpful for wildlife malaria and vet - taemias during chronic stage of infection [24, 25, 35]. erinary medicine researchers aiming identification of Importantly, the severity of disease caused by a given avian malaria infections. lineage of Plasmodium often varies markedly in different species of avian hosts, from absence of any clinical symp- toms to high mortality [4, 17, 19, 36–41]. Methods Because of broad vertebrate host specificity, the same Full-length papers with descriptions of new Plasmo- Plasmodium species can infect distantly related birds. In dium species published in peer-reviewed journals were other words, vertebrate host identity cannot be used as a considered. In all, 164 articles were reviewed, and 152 taxonomic feature during identification of avian malaria papers containing most representative information about Valkiūnas and Iezhova Malar J (2018) 17:212 Page 3 of 24 taxonomy of these parasites were incorporated in the of the cytoplasm in different haemosporidian species. References. While, this also depends on staining protocols, macro- Type and voucher preparation as well as images and microgametocytes can be readily distinguished of blood stages of avian Plasmodium parasites were in each haemosporidian species. This is not the case in obtained from the collections of Natural History other intracellular protists, whose gamonts and other Museum (London, UK), International Reference Centre intracellular blood stages do not show sexually dimorphic for Avian Haematozoa (Queensland Museum, Quens- features and all look similar under the light microscope land, Australia), the US National Parasite Collection (Fig. 1j–l). (National Museum of Natural History, Washington DC, Based on current taxonomy, four families of haemos- USA), Muséum National d’Histoire Naturelle (Paris, poridians can be recognized. These are Plasmodiidae, France), Grupo de Estudio Relación Parásito Hospedero, Haemoproteidae, Leucocytozoidae and Garniidae [1, 4, Universidad Nacional de Colombia (Bogotá, Colombia) 8, 30, 60, 61]. Malaria parasites are classified in the fam - and Nature Research Centre (Vilnius, Lithuania). All ily Plasmodiidae, which contains one genus Plasmodium. accessed preparations were studied. An Olympus BX61 When haemosporidians are found in blood films, Plas - light microscope (Olympus, Tokyo, Japan) equipped with modium parasites should be distinguished from species an Olympus DP70 digital camera and imaging software of related haemosporidians belonging to the families AnalySIS FIVE (Olympus Soft Imaging Solution GmbH, Garniidae, Haemoproteidae and Leucocytozoidae. The Münster, Germany) was used to examine preparations main distinctive features of parasites belonging to these and prepare illustrations. families are summarized in Table 1. A method of dichotomous key was applied for identifi - Blood stages of species of Plasmodium are particu- cation of Plasmodium species. This tool consists of steps larly similar to those of relatively rare haemosporidian divided it two alternative parts, which allow to determine parasites of the genera Fallisia and Garnia of the fam- the identity of a specimen due to a series of choices that ily Garniidae [8, 60–62]. Parasites of these three gen- lead the user to the correct name of a given specimen. era produce gametocytes and meronts (=schizonts) in The most difficult choices, which do not exclude ambi - blood cells (Fig. 1a–f ). However, species of Plasmodium guity, were accompanied with references to the corre- do not digest haemoglobin completely and accumulate sponding pictures, which illustrate meaning of the text residual pigment granules (hemozoin), which are refrac- information. This simplifies the comparison of diagnostic tory and readily visible in blood stages under light micro- features used in the keys. All parasite names in the keys scope (Fig. 1a–c). This is not true of species belonging to are accompanied with references to the original parasite the genera Fallisia and Garnia or other garniids, which descriptions and (or) reviews containing description and digest haemoglobin completely when they inhabit red (or) illustrations of corresponding species. blood cells and do not possess pigment granules in their blood stages (Fig. 1d–f ). Results When malaria parasites of the Plasmodium genus are Birds are often infected with different blood parasites reported in blood films, the next step is to distinguish belonging to same and different genera in the wild, and subgenera of this genus. The main characteristics of dif - various combinations of different parasite co-infections ferent subgenera are summarized in Table 2. often occur in same individual hosts. Haemosporidians When the subgenus of a malaria parasite has been (order Haemosporida) develop intracellularly, and they identified, the next step is the species identification using should be distinguished from other eukaryotic intracellu- the keys to species (Tables 3, 4, 5, 6). lar infections before identification of the parasite species identity. Haemosporidians can be readily distinguished Discussion from all other intracellular protists (species of Babesia, There are three main groups of obstacles, which a Isospora, Lankesterella, Haemogrerina, Hepatozoon, Tox- researcher usually faces during morphological identifica - oplasma) due to one particularly readily distinguishable tion of malaria parasites using microscopic examination feature. Mainly, gametocytes of all haemosporidians are of blood samples collected in the field. First, the quality characterized by sexually dimorphic features, which are of microscopic preparations is essential for correct par- readily distinguishable under the light microscope. Hae- asite identification, but often is insufficient due to thick mosporidian macrogametocytes possess compact nuclei blood films or artefacts of their drying, fixation, staining and bluish-stained cytoplasm, and the microgametocyte or storage. This precludes visualization of some impor - nuclei are diffuse and the cytoplasm stains paler than in tant features for species identification. It is essential to macrogametocytes (compare Fig. 1a, h with b, i). Some master these simple methods of traditional parasitology variation occurs in the size of nuclei and in the staining before sample collection, and this can be readily achieved Valkiūnas and Iezhova Malar J (2018) 17:212 Page 4 of 24 Fig. 1 Main morphological features of blood stages, which are used for identification of families of haemosporidian (Haemosporida) parasites (a–i). Mature gametocytes (a, b, g–i) and meronts (c–f) of Plasmodium (a–c), Garnia (d, e), Fallisia (f), Haemoproteus (g) and Leucocytozoon (h, i) parasites belonging to the families Plasmodiidae (a–c), Garniidae (d–f), Haemoproteidae (g) and Leucocytozoidae (h, i). Note presence of malarial pigment in species of Plasmodiidae (a–c) and Haemoproteidae (g) and its absence in species of Garniidae (d–f) and Leucocytozoidae (h, i). Macrogametocytes (a, g, h) and microgametocytes (b, i) are readily distinguishable due to presence of sexually dimorphic features. Common avian intracellular non-haemosporidian parasites (j–l) are shown for comparison with haemosporidians. These are Isospora (synonym Atoxoplasma) (j), Hepatozoon (k) and Babesia (l). Long simple arrows—nuclei of parasites. Simple arrowhead—pigment granules. Triangle arrowheads—developing merozoites. Long simple wide arrow—nucleolus. Simple wide arrowheads—host cell nuclei. Short simple wide arrow—cytoplasm of host cell. Scale bar = 10 µm. Explanations are given in the text Valkiūnas and Iezhova Malar J (2018) 17:212 Page 5 of 24 Table 1 Key to families of haemosporidian parasites Step Features and family 1 (4) Merogony takes place in blood cells (Fig. 1c–f ) 2 (3) Malarial pigment (hemozoin) is present in blood stages (Fig. 1a–c) …………………………………………… Plasmodiidae 3 (2) Malarial pigment (hemozoin) is absent from blood stages (Fig. 1d–f ) …………………………………………… Garniidae 4 (1) Merogony (Fig. 1c–f ) does not take place in blood cells. Only gametocytes (Fig. 1g–i) present in blood cells 5 (6) Malarial pigment (hemozoin) is present in blood stages (Fig. 1a, b, g) …………………………………………… Haemoproteidae 6 (5) Malarial pigment (hemozoin) is absent from blood stages (Fig. 1h, i) …………………………………………… Leucocytozoidae Main taxonomic features of families of the haemosporidian parasites  Merogony takes place in cells of fixed tissues and blood cells of vertebrate hosts. Malarial pigment (hemozoin) is present in erythrocytic meronts and gametocytes. Sexual process and sporogony of bird parasites take place in mosquitoes (Diptera: Culicidae) Merogony takes place in cells of fixed tissues and blood cells of vertebrate hosts. Malarial pigment (hemozoin) is absent at all stages. Vectors are still unknown Merogony takes place in cells of fixed tissues of vertebrate hosts. No merogony occurs in blood cells. Malarial pigment (hemozoin) is present in gametocytes. Sexual process and sporogony of bird parasites take place in louse flies (Hippoboscidae) and biting midges (Ceratopogonidae) Merogony takes place in cells of fixed tissues of vertebrate hosts. No merogony occurs in blood cells. Malarial pigment (hemozoin) is absent at all stages. Sexual process and sporogony take place in black flies (Simuliidae) and biting midges (Ceratopogonidae) in each laboratory using available protocols [1, 8, 63, 64]. research on Plasmodium species and recognition of new Second, Plasmodium species parasitaemia is often light malaria pathogens, for whose detection, detailed com- in natural infections in the wild. In other words, malaria parison with already described and genetically character- parasites might be reported in blood films, but not all ized parasites is needed. The development of molecular stages, which are needed for parasite species identifica - markers for diagnosis of disease agents is an important tion, are present. This might limit the use of the keys. task of current avian malariology (Table 7). Sampling of large number of birds (20–30 individuals) This study shows that 55 described species of avian belonging to the same species at a study site is often malaria parasites can be readily distinguished (Tables 3, helpful to detect relatively high parasitaemia of the same 4, 5, 6, 7). Among them, 12, 16, 22, 4 and 1 species belong pathogen and to access the full range of blood stages to subgenera Haemamoeba, Giovannolaia, Novyella, allowing parasite species identification. Third, co-infec - Huffia and Bennettinia, respectively. The great majority tions of Plasmodium species might occur, and requires of described avian Plasmodium species were reported some experience to distinguish between different patho - only in birds that live in tropical and subtropical coun- gens [45, 48, 56]. These obstacles strengthen the need for tries or in Holarctic migrants wintering in the same the development of molecular characterization in avian regions, indicating that transmission of these pathogens malaria diagnostics, which is still only available for 44% occurs mainly in countries with warm climates. Those of described parasite species, whose validity is obvious malaria parasites, which have adapted for transmission (Table 7). This is particularly timely for itemizing Plas - globally and have become cosmopolitan, are exceptions. modium species phylogenies, which currently are based Among these, Plasmodium relictum, Plasmodium elon- mainly on mitochondrial cytb gene sequences in avian gatum, Plasmodium circumflexum, Plasmodium matuti - malariology [5, 7, 23, 29, 33]. num and Plasmodium vaughani should be mentioned Molecular markers are sensitive for distinguishing dif- first of all [6, 8, 21, 23, 66–70]. These are invasive infec - ferent parasite species and their lineages, and they are tions, which are often virulent in non-adapted hosts, and essential for identification of cryptic Plasmodium species they are worth particular attention in bird health. . Molecular characterization is best developed for Among described avian Plasmodium parasites, species Novyella parasites (molecular markers are available for of Novyella are particularly diverse (Table 5). They rep - 59% of described species of this subgenus), and is weakest resent approximately 40% of all described avian malaria for Giovannolaia parasites (only two species or 12.5% of pathogens, and 78% of Plasmodium species, which were this subgenus have been characterized molecularly). Lack discovered during past 15 years. Novyella parasites are of molecular markers for many described malaria path- mainly pathogens of birds in countries of tropical and ogens [51, 53, 54, 56, 57, 59, 65] precludes biodiversity subtropical regions (Table 5). The Holarctic migrating Valkiūnas and Iezhova Malar J (2018) 17:212 Page 6 of 24 Table 2 Key to subgenera of Plasmodium parasites of birds Step Features and subgenus 1 (2) Exo-erythrocytic merogony takes place in cells of the haemopoietic system. Erythrocytic meronts develop in various immature red blood cells (Fig. 2i, k–p) …………………………………………… Huffia 2 (1) Exo-erythrocytic merogony does not takes place in cells of the haemopoietic system. Erythrocytic meronts do not develop in early immature red blood cells (Fig. 2i, k–p); mature and nearly mature erythrocytes are the main host cells (Figs. 2a–h, j; 3a–y) 3 (6) Roundish fully grown gametocytes (Fig. 4t–w) are present 4 (5) Size of fully grown gametocytes (Fig. 4u–w) and erythrocytic meronts (Fig. 1c) markedly exceed that of the nuclei of infected erythrocytes …………………………………………… Haemamoeba 5 (4) Size of fully grown gametocytes (Figs. 4a, 5m) and erythrocytic meronts (Fig. 3g, n) does not exceed that of the nuclei of infected erythrocytes …………………………………………… Bennettinia 6 (3) Roundish fully grown gametocytes (Fig. 4t–w) are absent. Elongate gametocytes (Fig. 4c–s) predominate 7 (8) Erythrocytic trophozoites (Fig. 3d) and growing meronts (Fig. 3x) contain plentiful cytoplasm. Size of fully grown erythrocytic meronts, which size markedly exceed that of the nuclei of infected erythrocytes (Figs. 2d, e, 3w, y), are present …………………………………………… Giovannolaia 8 (7) Erythrocytic trophozoites (Fig. 3a, b) and growing meronts (Fig. 3e–o) contain scanty cytoplasm. Size of fully grown erythrocytic meronts does not exceed or only slightly exceeds that of the nuclei of infected erythrocytes (Fig. 3p–r) …………………………………………… Novyella Main taxonomic characters of subgenera of avian malaria parasites  Exo-erythrocytic merogony takes place in cells of the haemopoietic system. Erythrocytic trophozoites and growing meronts (Fig. 2l–p) contain plentiful cytoplasm. Erythrocytic meronts develop in various immature red blood cells (Plasmodium huffi probably is an exception, but this needs confirmation). Fully grown erythrocytic meronts and gametocytes are variable both in form and size; elongate, roundish and irregularly shaped parasites might occur. Pedunculated oocysts are absent Exo-erythrocytic merogony takes place in cells of the reticuloendothelial system. Erythrocytic trophozoites (Fig. 3d) and growing meronts (Fig. 2a, b) contain plentiful cytoplasm. The size of fully grown erythrocytic meronts exceeds that of the nuclei of infected erythrocytes (Fig. 1c). Fully grown gametocytes are roundish, oval or of irregular form, and their size exceeds that of the nuclei of infected erythrocytes (Fig. 4t–x). Pedunculated oocysts are absent Exo-erythrocytic merogony takes place in cells of the reticuloendothelial system. Erythrocytic trophozoites and growing meronts contain scanty cytoplasm (Fig. 3g). Growing erythrocytic meronts are nucleophilic. The size of fully grown erythrocytic meronts does not exceed that of the nuclei of infected erythrocytes (Fig. 3g, s). Fully grown gametocytes are roundish, oval, of irregular form, sometimes oval-elongated; their size does not exceed that of the nuclei of infected erythrocytes (Fig. 5m). Pedunculated oocysts are present. Subgenus Bennettinia contains only one species, Plasmodium juxtanucleare  Exo-erythrocytic merogony takes place in cells of the reticuloendothelial system. Erythrocytic trophozoites (Fig. 3d) and growing meronts (Fig. 3x) contain plentiful cytoplasm. The size of fully grown erythrocytic meronts exceeds that of the nuclei of infected erythrocytes (Figs. 2d, e; 3w, y). Fully grown gametocytes are elongated (Figs. 4c–s; 5i, k, o). Pedunculated oocysts are absent Exo-erythrocytic merogony takes place in cells of the reticuloendothelial system. Erythrocytic trophozoites (Fig. 3a, b) and growing meronts (Fig. 3e–j) contain scanty cytoplasm. The size of fully grown erythrocytic meronts does not exceed or only slightly exceeds that of the nuclei of infected erythrocytes occasionally (Fig. 3p–r). Fully grown gametocytes are elongated (Fig. 4c–r). Pedunculated oocysts are absent birds gain Novyella infections in their wintering grounds or of low prevalence in areas with cold climates located and transport them to their breeding grounds where they close to the Polar Circles [1, 8, 18, 19, 58, 68]. are normally not transmitted [8, 71–73]. Factors prevent- Limited available experimental information indicates ing spread of Novyella infections globally are unclear. that some Novyella species (P. ashfordi, P. rouxi) may Novyella species are the most poorly studied group of cause severe and even lethal malaria in some birds due to avian malaria pathogens, with nearly no information blood pathology [1, 8, 74, 75], but the complete mecha- available about exo-erythrocytic development, virulence, nism of their pathogenicity remains unresolved, mainly sporogony and vectors for the great majority [1, 4, 8, due to lack of information about exo-erythrocytic devel- 72]. A few Novyella parasites (P. vaughani, Plasmodium opment . Investigation of life cycles and virulence of rouxi, Plasmodium homopolare) are actively transmitted infections caused by Novyella species is an important in countries with temperate climates, but they are absent task in current avian malaria research. Valkiūnas and Iezhova Malar J (2018) 17:212 Page 7 of 24 Many species of Plasmodium inhabit numerous species A list of the Plasmodium species names of unknown of birds and use mosquitoes of different genera for trans - taxonomic position (incertae sedis) and also the names mission [1, 8, 9, 11]. Within this spectrum of hosts and of species of doubtful identity, which require further vectors, the same parasite species might exhibit diverse investigation (species inquirenda), is given in Table 9. All morphological forms and strain varieties. Because of these parasite descriptions are insufficiently complete these morphological variants, it has been conventional in and were not accompanied with molecular characteri- old avian malaria research (approximately between 1927 zation. Taxonomic status of the majority of these names and 1995) that any new Plasmodium species description was justified in . Twenty names of Plasmodium para- should only be accepted if supported by a comprehensive sites were added to this list and their taxonomic status package of taxonomic features, which not only included was explained (Table 9). The majority of these parasite the full range of blood stages, but also data on the verte- descriptions are based on preparations with co-infections brate host specificity, periodicity of erythrocytic merog - of several Plasmodium parasites belonging to same and ony, tissue merogony, vectors and patterns of sporogonic (or) different genera. This raises a question if all blood development. It is not surprising that recent molecular stages reported in the original descriptions truly belong studies supported the validity of the old Plasmodium to corresponding species. species descriptions, which were detailed and precise Additionally, in many of such parasite descriptions, (Table 7). Application of molecular diagnostic tools in gametocytes were not described, but this stage is essen- studies of avian haemosporidian parasites [29, 69, 76, 77] tial for the identification of some Plasmodium species opened new opportunities to distinguish haemosporid- (Tables 3, 4, 5, 6, Figs. 4, 5). It is important to note that ian parasites based on their unique DNA sequences. This the descriptions of many Plasmodium parasites, which stimulated biodiversity research of wildlife Plasmodium were incorporated in Table 9 and published during past parasites, particularly because the molecular characteri- 15 years, contain some information about their blood zation, which was done in parallel with morphological stages. Additionally, the type material was designated in description of blood stages, made each parasite species many descriptions, but usually is insufficient for practi - detection readily repeatable at all stages of life cycle cal use and distinguishing parasites at the species level, (Table 7). particularly because (1) the type preparations contain co- A list of synonymous names of avian Plasmodium spe- infections and (2) single cells (meronts) were designated cies and the justification of the nomenclature status of as holotypes. Single cells usually do not reflect entire these names are given in Table 8. The majority of these morphological diversity of malaria parasites, so deposi- parasite descriptions are insufficiently complete and were tion of parahapantotype material is preferable in wildlife not accompanied with molecular characterization. Due haemosporidian research [35, 49, 58, 78]. Validation of to the huge genetic diversity of avian malaria pathogens some names listed in Table 9 is possible in the future, but and numerous genetic lineages reported in birds, some it requires additional research, preferably based on new of these names might be validated in the future, and they samples from the same avian hosts and type localities. represent a reserve for future taxonomic work. However, Invalid Plasmodium parasite names (nomen nudum) available descriptions of these parasites do not provide are listed in Table 9. These names were not accompanied sufficient information to readily distinguish them from with descriptions so have no status in nomenclature. The parasites, whose validity is well established (Tables 3, 4, 5, names of this category can be used as a reserve for new 6). For clearness of scientific texts, it is preferable to avoid parasite descriptions in the future, but it is preferable not use of the synonymous names before additional data on to use them to avoid taxonomic confusion . their validity are available. Reports of parasite lineages The subgenus Papernaia was created for Novyella-like and GenBank accessions of their DNA sequences in pub- avian malaria parasites, whose erythrocytic meronts do lications would be helpful to specify Plasmodium species not possess globules (Fig. 3f, h–l), structures of unclear identity in the future. origin and function [79, 80]. The feature of the presence or absence of such globules is used in distinguishing Valkiūnas and Iezhova Malar J (2018) 17:212 Page 8 of 24 Table 3 Key to the Haemamoeba species Step Features and species 1 (16) Roundish or oval pigment granules predominate in gametocytes (Fig. 4u–x). Elongate rod-like in form pigment granules (Fig. 5n) are absent, but single slightly elongate pigment granules might occur occasionally 2 (22) A residual body (Fig. 5s) is absent in mature erythrocytic meronts. Of oval-elongate form gametocytes, which are over 10 µm in length (Fig. 5i), are present 3 (17) Large (≥ 1 µm in diameter) vacuoles (Figs. 2g, h, 5u) are absent from growing erythrocytic meronts. Markedly vacuolated erythrocytic meronts (Fig. 2f–h) are absent 4 (9) Maximum number of merozoites in mature (Figs. 2e, j, 3n–r, y, 5r, s) erythrocytic meronts is ≤ 12 5 (8) Maturing and mature erythrocytic meronts enlarge infected erythrocytes < ½ in area in comparison to uninfected erythrocytes (compare infected and unin- fected erythrocytes in Fig. 2g, h, see also Fig. 5s); numerous mature meronts adhere to erythrocyte nuclei (Fig. 2j) 6 (7) Merozoites locate haphazardly in mature meronts (Fig. 1c). Residuum cytoplasm (Fig. 5s) is invisible in mature meronts, and merozoites never appear to have connections to the residuum cytoplasm ……………………………………………. P. subpraecox [1, 8, 82] 7 (6) Nuclei locate on periphery of maturing and mature meronts (Fig. 5r). Residuum cytoplasm is visible and locates centrally in maturing meronts (Fig. 5r). Maturing merozoites have connections to the residuum cytoplasm, and these connections look like small wisps of cytoplasm extending towards merozoites (Fig. 5r) ……………………………………………. P. parvulum  8 (5) Maturing and mature meronts enlarge infected erythrocytes over ½ in area in comparison to uninfected erythrocytes (Fig. 5p). The majority of maturing meronts are rounded in shape, they locate away from erythrocyte nuclei, which are markedly displaced toward erythrocyte envelope from earliest stages of meronts development (Fig. 5p) ……………………………………………. P. caloti  9 (4) Maximum number of merozoites in mature erythrocytic meronts is > 12. Mature meronts and gametocytes are large (size is significantly greater than erythrocyte nuclei); they occupy > 1/2 of the cytoplasm in infected erythrocytes (Figs. 2j; 4u–x) 10 (23) Pigment granules in gametocytes do not tend to be clumped in a spot, which is usually located near a margin of the parasite (Fig. 4t, w). If present occasionally, such position of pigment granules does not predominate in mature gametocytes 11 (10) Pigment granules in mature gametocytes show markedly different patterns of position in the cytoplasm; they often are randomly scattered (Figs. 1b, 5v), but also might be variously grouped (Fig. 5t) and even aggregated in solid masses (Fig. 4u, v, x) 12 (15) Largest fully grown gametocytes can occupy all available cytoplasmic space in infected erythrocytes (Fig. 5x, y). Length of the largest gametocytes exceed 10 µm 13 (14) Development in the blood is asynchronous, with all blood stages present in circulation simultaneously. Periodicity of erythrocytic merogony is 36 h; Specific parasite of domestic chicken. Passeriform birds are resistant. In the nature, transmission does not occur outside the Oriental zoogeographical region ……………………………………………. P. gallinaceum [1, 8, 83] 14 (15) Development in the blood is synchronous, with not all blood stages present in circulation simultaneously. Periodicity of erythrocytic merogony is 24 h. Domestic chicken was reported to be resistant. In the nature, transmission occurs outside the Oriental zoogeographical region ……………………………………………. P. coturnixi [8, 84] 15 (12) Largest fully-grown gametocytes do not occupy all available cytoplasmic space in infected erythrocytes; a small non-occupied space is usually visible in infected erythrocytes (Fig. 4u–x). Length of the largest gametocytes does not exceed 10 µm. Domestic chicken is resistant. Development in the blood is asynchronous, with all blood stages (trophozoites, growing and mature meronts as well gametocytes) present in blood films simultaneously. Periodicity of erythrocytic merogony is 36 h ……………………………………………. P. relictum [8, 26, 85] 16 (1) Pigment granules in gametocytes are roundish, oval and elongate rod-like (Fig. 5n). Rod-like pigment granules are common and might predominate in microga- metocytes (Fig. 5n), but they are less common and often do not predominate in macrogametocytes ……………………………………………. P. cathemerium [1, 8, 86] 17 (3) Large (≥ 1 µm in diameter) vacuoles (Figs. 2h; 5u) are common in erythrocytic meronts 18 (21) One or several large vacuoles, which do not exceed 2 µm in diameter, are often present in growing erythrocytic meronts. Markedly vacuolated erythrocytic meronts are common (Fig. 2f–h). Pigment granules do not gather around these vacuoles. Trophozoites lack large (> 1 µm in diameter) vacuoles. Lobulated in form gametocytes (Fig. 4x) are absent or develop only occasionally 19 (20) Vacuoles are absent or occur occasionally in erythrocytic trophozoites. Pigment granules in fully grown gametocytes distinctly vary in size, and small (< 0.5 µm) and medium (0.5–1.0 µm) size granules occur simultaneously (Fig. 5v). The medium-size pigment granules are common (Fig. 5v). Phanerozoites do not develop in brain of domestic canaries ……………………………………………. P. giovannolai [1, 8, 87] 20 (19) Vacuoles often present in erythrocytic trophozoites. Pigment granules in fully grown gametocytes are more or less similar in size, usually they are small (< 0.5 µm) (Fig. 5t). Medium-size (0.5–1.0 µm) pigment granules (Fig. 5v) might occur, but are not characteristic. Phanerozoites develop in brain of domestic canaries ……………………………………………. P. matutinum [1, 8, 66, 88] 21 (19) Each advanced trophozoites possess one large (> 1 µm in diameter) roundish centrally located vacuole. One large (> 2 µm in diameter) vacuole is present in growing erythrocytic meronts (Fig. 5u). Pigment granules gather around this vacuole. Lobulated in form gametocytes (Fig. 4x) are common ……………………………………………. P. tejerai [8, 50, 89] Valkiūnas and Iezhova Malar J (2018) 17:212 Page 9 of 24 Table 3 (continued) Step Features and species 22 (2) A residual body (Fig. 5s) is present in mature erythrocytic meronts. Of oval-elongate form gametocytes, which are over 10 µm in length (Fig. 5i), are present. Growing erythrocytic meronts often possess vacuoles (Fig. 2g, h) ……………………………………………. P. griffithsi [1, 8] 23 (10) Pigment granules in gametocytes clearly tend to be clumped in a spot, which is located near a margin of the parasite (Fig. 4t, w). Such position of pigment granules predominates in mature gametocytes. Pigment granules can be aggregated into a solid mass of pigment, which also usually locates near a margin of the parasite ……………………………………………. P. lutzi [8, 90, 91] Plasmodium caloti was described from the Eurasian skylarks Alauda arvensis co-infected with several other Plasmodium species, and this races a question if all blood stages (particularly gametocytes), which were reported in the original description , truly belong to this parasite. However, because of (1) the marked influence on host cell (marked enlargement of infected erythrocytes and displacement of their nuclei) and (2) the relatively regular rounded form and smooth margins of mature meronts (Fig. 5p), which produce small number of merozoites < 10), this parasite is morphologically unique and can be distinguished from other Haemamoeba species. The original description is fragmentary , and re-description of this parasite is needed Table 4 Key to Giovannolaia species Step Features and species 1 (16) Elongate meronts, which grow laterally to nuclei of infected erythrocytes (Figs. 2c–e; 3w–y), predominate 2 (3) Cytoplasm of gametocytes (especially macrogametocytes) is highly vacuolated (Fig. 4o). Large (> 1.5 µm in diameter) vacuoles are present in some macrogametocytes ……………………………………………… P. fallax [1, 8, 92] 3 (2) Cytoplasm of gametocytes is not highly vacuolated; if vacuoles are present in macrogametocytes, they are few and of small size (< 1 µm in diameter) (Fig. 4p–s) 4 (5) Pigment granules in the majority of erythrocytic meronts are aggregated into large (> 1.5 µm in length) clumps, which usually locate at one end of elongate meronts (Fig. 5f) ……………………………………………… P. anasum [1, 8, 93] 5 (4) Pigment granules in the majority of erythrocytic meronts are not aggregated into large (> 1.5 µm in length) clumps, which usually locate at one end of elongate meronts (Fig. 5f ). Location of pigment granules in erythrocytic meronts is markedly variable 6 (7) Nuclei tend to lean to one end in the majority of growing erythrocytic meronts (Fig. 5d) ……………………………………………… P. leanucleus [8, 94] 7 (6) Nuclei do not tend to lean to one end in the majority of growing erythrocytic meronts (Fig. 5d). Position of nuclei in developing meronts is markedly variable (Fig. 3u, x) 8 (11) Average number of merozoites in mature meronts is < 12 9 (10) Fully grown erythrocytic meronts and gametocytes are thin slender cells, they do not displace the nuclei of infected erythrocytes and usually do not adhere to the nuclei (Fig. 5e) ……………………………………………… P. gundersi [1, 8, 95] 10 (9) Fully grown erythrocytic meronts (Fig. 3w) and gametocytes (Fig. 5k) are broad cells, which width is equal to the width of erythrocyte nuclei or is greater; both mature meronts and gametocytes displace the nuclei of infected erythrocytes laterally and often adhere to the nuclei (Figs. 3w; 5k) ……………………………………………… P. octamerium [8, 96] 11 (8) Average number of merozoites in mature meronts is ≥ 12 12 (15) Gametocytes and meronts grow around nuclei of erythrocytes (Figs. 2c, d; 4r, s). Fully grown erythrocytic meronts and gametocytes usually only slightly (if at all) influence infected erythrocytes and do not displace or only slightly displace nuclei of erythrocytes laterally (Figs. 2d, e; 4s). Infected erythrocytes usually do not become rounded (Fig. 5x) 13 (14) Fully-grown erythrocytic meronts (Fig. 2d, e) and gametocytes (Fig. 4s) markedly (often nearly completely or completely) encircle nuclei of infected erythrocytes; completely circumnuclear mature meronts and gametocytes frequently develop, but their occurrence depends of stage of parasitemia, so they might be not always seen in blood films ……………………………………………… P. circumflexum [1, 8, 97] , P. homocircumflexum  14 (13) Fully grown erythrocytic meronts never assume circumnuclear form (Fig. 3w, y). Gametocytes nearly completely (Fig. 4r) or completely (Fig. 4s) encircle nuclei of infected erythrocytes; completely circumnuclear mature gametocytes develop, but usually are rare. Advanced trophozoites and young meronts often possess large (> 1 µm in diameter) vacuoles (Fig. 5j) ……………………………………………… P. lophurae [1, 8, 98] Valkiūnas and Iezhova Malar J (2018) 17:212 Page 10 of 24 Table 4 (continued) Step Features and species 15 (12) Gametocytes and meronts start to grow around nuclei of erythrocytes However, fully grown meronts markedly displace nuclei of erythrocytes and assume various irregular forms; they often roundish or close to roundish in shape (Fig. 5w), markedly displace the nuclei of infected erythrocytes and can occupy all available cytoplasmic space in the erythrocytes (Fig. 5w). Fully grown gametocytes markedly deform infected erythrocytes, which become rounded (Fig. 5x, y) ……………………………………………… P. gabaldoni [8, 99] 16 (1) Elongate erythrocytic meronts, which grow laterally to nuclei of infected erythrocytes (Figs. 2c–e; 3w–y), are absent or appear only occasion- ally; they never predominate. The majority of fully grown meronts are of roundish, oval or irregular form; they do not take or take only occasionally the lateral position to nuclei of erythrocytes (Fig. 5q–s) 17 (24) Large (> 1.5 µm in diameter) vacuoles (Fig. 5g) absent from gametocytes. If small vacuoles are present in gametocytes, pigment granules do not gather around vacuoles 18 (25) Fully grown gametocytes do not tend to lie obliquely in infected erythrocytes (Fig. 5i, o), and they do not displace the nuclei towards one pole of the erythrocytes 19 (26) Growing erythrocytic meronts do not produce long (> 2 µm in length) tail-like or finger-like outgrowths (Fig. 5c) 20 (21) Erythrocytic meronts take a polar or subpolar position in infected erythrocytes, and their influence on infected erythrocytes is usually not pronounced (Fig. 3r) ……………………………………………… P. polare [1, 8, 100] 21 (20) Erythrocytic meronts can be seen anywhere in infected erythrocytes including a lateral, subpolar and polar position. If meronts take a polar or subpolar position in the erythrocytes, they markedly influence the host cells causing their deformation and (or) displacement of their nuclei 22 (23) Maximum number of merozoites in mature meronts > 10. Size of pigment granules in macro- and microgametocytes is clearly different ……………………………………………… P. pinottii [1, 8, 101] 23 (22) Maximum number of merozoites in mature meronts < 10. Size of pigment granules in macro- and microgametocytes is similar ……………………………………………… P. garnhami [1, 8, 102] 24 (17) Large (> 1.5 µm in diameter) vacuoles (Fig. 5g) develop in many macrogametocytes. Pigment granules often gather around these vacuoles ……………………………………………… P. formosanum [1, 8, 103] 25 (18) Fully grown gametocytes tend to lie obliquely in infected erythrocytes, and they displace the nuclei towards one pole of the erythrocytes (Fig. 5o) ……………………………………………… P. durae [1, 8, 104] 26 (19) Growing erythrocytic meronts often produce long (> 2 µm in length) tail-like or finger-like outgrowths (Fig. 5c) 27 (28) Nuclei in mature erythrocytic meronts are usually arranged as fans (Fig. 3v), rosettes (Fig. 5r), or more or less pronounced rows (Fig. 3w). Infected erythrocytes with segmented mature meronts are often rounded (Fig. 5r). Fully grown gametocytes do not fill erythrocytes up to their poles (Fig. 4h) ……………………………………………… P. pedioecetae [1, 8, 105, 106] 28 (27) Nuclei in mature erythrocytic meronts are usually located randomly (Fig. 3q) and they only occasionally can be arranged as rosettes. Infected erythrocytes with segmented mature meronts are not rounded (Fig. 3w). Fully grown gametocytes fill erythrocytes up to their poles (Fig. 5k) ……………………………………………… P. hegneri [8, 93] Based on available information, P. circumflexum and P. homocircumflexum are cryptic species, which cannot be distinguished using morphological features of their blood stages . Cytochrome b sequences can be used to distinguish these infections (see Table 7) Valkiūnas and Iezhova Malar J (2018) 17:212 Page 11 of 24 Table 5 Key to the Novyella species Step Features and species 1 (19) Maximum number of merozoites in erythrocytic meronts > 4 2 (41) Maturing erythrocytic meronts, which displace host-cell nuclei, assume a fan-like shape and possess elongate nuclei (Fig. 3v), are absent 3 (26) Erythrocytic meronts, which lie free in the cytoplasm of host cell and do not touch the nuclei of infected erythrocytes (Fig. 3e–l, o), are present 4 (32) Trophozoites and binuclear meronts (Fig. 3e, f ) do not produce clearly defined long outgrowths (Fig. 5a); if ameboid outgrowths are present, they do not exceed the main body of the trophozoites in length 5 (42) Ends of growing macrogametocytes are similar in width (Fig. 4c–e, g–r) 6 (9) Number of merozoites in mature erythrocytic meronts is relatively stable. Over 90% of the mature meronts contain 6 merozoites 7 (8) Macrogametocyte nuclei are terminal in position (Fig. 4g). Refractive globules (Fig. 3h) are present in erythrocytic meronts ……………………………………………. P. parahexamerium  8 (7) Macrogametocyte nuclei are central or subcentral in position (Fig. 4o). Refractive globules are absent from erythrocytic meronts (Fig. 3p) ……………………………………………. P. hexamerium [1, 8, 55, 107] 9 (6) Number of merozoites in mature erythrocytic meronts is variable 10 (37) Growing and mature meronts assume various positions to the erythrocyte nuclei; they can be found in polar, sub-polar and lateral position in relation to the host cell nuclei 11 (38) Binuclear erythrocytic meronts do not possess large (of size, which is similar to nuclei of the meronts), centrally located vacuoles (Fig. 3e). Macro- and microga- metocytes are of similar shape, they assume similar positions in erythrocytes (Fig. 4g, h) 12 (39) Gametocytes do not possess refractive globules 13 (16) Erythrocytic meronts possess globules in natural infections (Fig. 3f, h–j) 14 (15) The majority of trophozoites as well as developing and mature erythrocytic meronts possess one of circular shape, prominent (on average 0.5 µm in area) pig- ment granule (Fig. 3q). Fan-like in shape mature meronts predominate ……………………………………………. P. unalis  15 (14) The majority of trophozoites, developing and mature erythrocytic meronts possess 1–4 (usually 2–3) small (< 0.5 µm in area), of different size pigment granules (Fig. 3c, h). Fan-like in shape mature meronts (Fig. 3o) are uncommon ……………………………………………. P. vaughani [1, 8, 49, 55, 108] 16 (13) Erythrocytic meronts do not possess globules in natural infections (Fig. 3g, n, o, r, s) 17 (18) Fan-like mature meronts containing 7–8 merozoites are common (Fig. 3o); pigment granules in gametocytes are clumped together into a prominent group, which is predominantly of terminal position in the gametocytes (Fig. 4e, h) ……………………………………………. P. ashfordi  18 (17) Fan-like mature meronts containing 7–8 merozoites are absent; pigment granules in gametocytes are scattered or clumped, but position of these clumps is irregular (never predominantly terminal) in the gametocytes ……………………………………………. P. forresteri [8, 109] 19 (1) Maximum number of merozoites in erythrocytic meronts is 4 20 (21) Erythrocytic meronts do not possess globules in natural infections (Fig. 3g) ……………………………………………. P. bertii [8, 110] 21 (20) Erythrocytic meronts possess globules in natural infections (Fig. 3f, h–l) 22 (23) One small (< 0.5 µm in diameter) refractive globule present in the majority of meronts (Fig. 3f, h, j). Blue non-refractive globules (Fig. 3k, l) are absent from mer- onts. The cytoplasm in gametocytes is more or less homogenous, but never is granular-like (Fig. 4c) or globular-like (Fig. 4l, m) in appearance ……………………………………………. P. rouxi [1, 8, 52, 111] 23 (22) Refractive globules (Fig. 3f, h–j) are absent from meronts. One blue non-refractive globule present in each advanced trophozoite (Fig. 3b), growing and mature meront (Fig. 3k, l). The cytoplasm in gametocytes is granular-like (Fig. 4c) or globular-like in appearance (Fig. 4l, m) 24 (25) One large (size similar to parasite nuclei or greater) blue non-refractive globule present in each advanced trophozoite (Fig. 3b), growing and mature meront (Fig. 3l). The cytoplasm in macro- and microgametocytes is markedly globular-like in appearance (Fig. 4l, m). Average number of pigment granules in macro- and microgametocytes is close to 10 ……………………………………………. P. megaglobularis  25 (24) One small (size smaller than parasite nuclei) blue non-refractive globule present in each advanced trophozoite, growing and mature meront (Fig. 3k). The cytoplasm in gametocytes is markedly granular in appearance, which is better visible in macrogametocytes (Fig. 4c); globular-like appearance of the cytoplasm (Fig. 4l, m) is not characteristic. Average number of pigment granules in macro- and microgametocytes is close to 5 ……………………………………………. P. globularis  26 (3) Erythrocytic meronts, which lie free in the cytoplasm of host cell and do not touch the nuclei of infected erythrocytes (Fig. 3e–l, o), are absent. Erythrocytic meronts are strictly nucleophilic (Figs. 3n, s; 5q) 27 (40) Both meronts (Fig. 3n, p, s, t) and gametocytes (Fig. 4a, d) are strictly nucleophilic 28 (29) Large (≥ 1 mµ in length) pigment granules are present in gametocytes (Fig. 4d); both mature gametocytes (Fig. 4d) and meronts (Fig. 3n, t) do not displace nuclei of erythrocytes ……………………………………………. P. delichoni  29 (30) Large (≥ 1 mµ) pigment granules (Fig. 4d) are absent from gametocytes; mature meronts (Fig. 5q) or mature gametocytes (Fig. 5k) displace nuclei of erythrocytes Valkiūnas and Iezhova Malar J (2018) 17:212 Page 12 of 24 Table 5 (continued) Step Features and species 30 (31) Fully-grown gametocytes do not fill erythrocytes up to their poles (Fig. 4h). Fully grown meronts displace nuclei of erythrocytes (Fig. 5q). Fully-grown gameto - cytes do not displace the nuclei of infected erythrocytes (Fig. 4h) ……………………………………………. P. nucleophilum [1, 8, 112–114] 31 (30) Fully grown gametocytes fill erythrocytes up to their poles (Fig. 5k). Fully grown erythrocytic meronts do not displace or only slightly displace the nuclei of infected erythrocytes (Fig. 3t). Fully-grown gametocytes markedly displace the nuclei of infected erythrocytes laterally (Fig. 5k) ……………………………………………. P. paranucleophilum [8, 113] 32 (4) Trophozoites and (or) binuclear meronts often produce clearly defined long outgrowths (Fig. 5a); the outgrowths exceed the main body of the trophozoites in length 33 (34) One or two refractive globules present in advanced trophozoites and developing and mature meronts. Each globule has a clear rim at its periphery (Fig. 3i); macrogametocytes are markedly vacuolated in appearance (Fig. 4n) ……………………………………………. P. multivacuolaris  34 (33) Globules are absent from trophozoites and meronts, but vacuoles might be present 35 (36) A large (> 1 µm in length) vacuole is present in trophozoites (Fig. 5a). Number of merozoites in erythrocytic meronts is relatively stable; approximately 95% of the meronts contain five merozoites. Pigment granules in gametocytes either randomly scattered throughout the cytoplasm or clumped into several small groups ……………………………………………. P. kempi [8, 115] 36 (35) A large (> 1 µm in length) vacuole (Fig. 5a) is absent from trophozoites, but a small (< 1 µm in diameter) vacuole present occasionally. Number of merozoites in erythrocytic meronts is variable, but most often is equal to 8. Pigment granules in gametocytes frequently are clumped in a focus near one end of the gameto- cytes (Fig. 4h) ……………………………………………. P. columbae [1, 8, 116] 37 (10) Growing and mature meronts are strictly of polar or subpolar position to the erythrocyte nuclei, and they usually do not adhere to the nuclei (Fig. 3q). Refractive globules present in grooving meronts, but often are invisible in mature meronts ……………………………………………. P. homopolare  38 (11) Binuclear erythrocytic meronts often possess one large (size similar to nuclei of the meronts), centrally located vacuole (Fig. 3e). In binuclear meronts, nuclei locate asymmetrically in relation to the vacuole (Fig. 3e). Shape of macro- and microgametocytes is different: microgametocytes are elongate (Fig. 4j), but mac- rogametocytes are not, and they are more or less roundish, lobulated or irregular in form parasites (Fig. 4b). Macrogametocytes usually take polar or subpolar position to nuclei of erythrocytes (Fig. 4b), and microgametocytes locate laterally to the nuclei of erythrocytes (Fig. 4j). Trophozoites possess one clear vacuole (Fig. 3a), which maintain in fully grown meronts (Fig. 3m) ……………………………………………. P. lucens  39 (12) Gametocytes possess refractive globules (Fig. 5h). Refractive circular globules present in macro- and microgametocytes ……………………………………………. P. accipiteris  40 (27) Meronts are strictly nucleophilic (Fig. 3p), but gametocytes are not (Fig. 4r). The majority of gametocytes do not adhere to nuclei of infected erythrocytes. Mature gametocytes markedly enclose nuclei of erythrocytes by their ends (Fig. 4r) ……………………………………………. P. homonucleophilum  41 (2) Maturing erythrocytic meronts, which displace host-cell nuclei, assume fan-like shape and possess elongate nuclei (Fig. 3v), are present. Number of nuclei in maturing fan-like meronts is about 10–12 ……………………………………………. P. valkiunasi  42 (5) Ends of growing macrogametocytes are markedly different in width ……………………………………………. P. dissanaikei [1, 8, 118] Plasmodium valkiunasi was described from Eurasian magpies Pica pica co-infected with several other Plasmodium species, and this races a question if all blood stages (particularly gametocytes), which were reported in the original description , truly belong to this species. However, this parasite is morphologically unique and can be distinguished from other Novyella species because of unique shape of its maturing meronts (Fig. 3v), which are large, develop in mature erythrocytes, have a regular fan-like form and possess numerous (about 12) peripherally located elongate nuclei. The original description is fragmentary , and re-description of this parasite is needed some species of malaria parasites belonging to subgenus artificial passages in unusual avian hosts. This strain was Novyella during natural infections (Table 5). It is inter- originally isolated from the Common cuckoo Cuculus esting to note that experimental studies with a Plasmo- canorus (Cuculiformes), and it did not possessed glob- dium ashfordi (pGRW2) strain, which normally do not ules in erythrocytic meronts  in the cuckoo or during possess globules in erythrocytic meronts, show that the the first passage in the Eurasian siskin Carduelis spinus. globules appeared in this parasite’s meronts after several However, the globules appeared in the meronts of the Valkiūnas and Iezhova Malar J (2018) 17:212 Page 13 of 24 Table 6 Key to Huffia species Step Features and species 1 (2) Development and maturation of gametocytes occurs in various immature red blood cells, including erythroblasts (Fig. 4y). The outline of nuclei in growing erythrocytic meronts (Fig. 2m, n) as well as growing and mature gametocytes (Fig. 4y) is smooth, and boundaries between nuclei and the cytoplasm are strictly distinct (Figs. 2m, n; 4y) ………………………………………………… P. polymorphum  2 (1) Development and maturation of gametocytes occurs only in mature or nearly mature red blood cells (Fig. 4p, q); gametocytes do not develop in erythroblasts. The outline of nuclei in growing erythrocytic meronts (Fig. 2l, o, p) as well as growing and mature gametocytes (Fig. 4p, q) is markedly variable, predominantly not smooth, and boundaries between the nuclei and the cytoplasm are often poorly distinct (Figs. 2o, p; 4p, q), particularly in the growing parasites 3 (6) In peripheral blood, trophozoites and erythrocytic meronts develop mainly in young red blood cells. Maximum number of merozoites in erythrocytic meronts is less than 20 4 (5) Elongated erythrocytic merozoites are present (Fig. 2k). Fully grown gametocytes are slender; they do not displace or only slightly displace the nuclei of infected erythrocytes laterally (Fig. 4p, q). Maximum width of fully grown gametocytes is equal or less than the width of nuclei of host cells (Fig. 4p, q) ………………………………………………… P. elongatum [1, 8, 23, 119, 120] 5 (4) Elongated erythrocytic merozoites (Fig. 2k) are absent. Fully grown gametocytes are broad; they markedly displace the nuclei of infected erythrocytes laterally and can fill the poles of infected erythrocytes completely (Fig. 5l). Maximum width of fully grown gametocytes is greater than the width of nuclei of host cells (Fig. 5l) ………………………………………………… P. hermani [8, 120, 121] 6 (3) In peripheral blood, trophozoites and erythrocytic meronts develop mainly in mature red blood cells. Maximum number of merozoites in erythrocytic meronts is greater than 20 ………………………………………………… P. huffi [1, 8, 122] Table 7 Mitochondrial cytochrome b sequences, which have been developed for molecular detection and identification (barcoding) of avian Plasmodium parasites a b Parasite subgenus and species GenBank accession and lineage code (in parentheses) References Haemamoeba P. caloti Not available Not available P. cathemerium AY377128 (pSEIAUR01)  P. coturnixi Not available Not available P. gallinaceum AY099029 (pGALLUS01)  P. giovannolai Not available Not available P. griffithsi Not available Not available P. lutzi KC138226 (pTFUS05)  P. matutinum KY287235 (pLINN1)  P. parvulum Not available Not available P. subpraecox Not available Not available P. relictum AF495571 (pSGS1), AY831748 (pGRW11), AY099041 (pGRW4), KC342644 (pLZ- [67, 71, 117] FUS01), MG724747 (pPHCOL01) P. tejerai JX272844 (pSPMAG01)  Giovannolaia P. anasum Not available Not available P. circumflexum AF495576 (pTURDUS1)  P. durae Not available Not available P. fallax Not available Not available P. formosanum Not available Not available P. gabaldoni Not available Not available P. garnhami Not available Not available P. gundersi Not available Not available P. hegneri Not available Not available P. homocircumflexum KC884250 (pCOLL4)  Valkiūnas and Iezhova Malar J (2018) 17:212 Page 14 of 24 Table 7 (continued) a b Parasite subgenus and species GenBank accession and lineage code (in parentheses) References P. leanucleus Not available Not available P. lophurae Not available Not available P. polare Not available Not available P. octamerium Not available Not available P. pedioecetae Not available Not available P. pinottii Not available Not available Novyella P. accipiteris Not available Not available P. ashfordi AF254962 (pGRW2)  P. bertii Not available Not available P. columbae Not available Not available P. delichoni KU529943 (pCOLL6)  P. dissanaikei Not available Not available P. forresteri Not available Not available P. globularis EU770151 (pANLA1)  P. hexamerium Not available Not available P. homonucleophilum KC342643 (pSW2)  P. homopolare KJ482708 (pSOSP CA 3P)  P. kempi Not available Not available P. lucens FJ389156 (pCYOL2)  P. megaglobularis EU770152 (pCYOL1)  P. multivacuolaris FJ389157 (pANLA2)  P. nucleophilum JX467689 (pEG01)  P. parahexamerium FJ389155 (pALDI1)  P. paranucleophilum Not available Not available P. rouxi HM146901 (pPADOM16)  P. unalis KC771247 (pTFUS06)  P. valkiunasi Not available Not available P. vaughani DQ847271 (pSYAT05)  Bennettinia P. juxtanucleare AB250415 (pGALLUS02)  Huffia P. elongatum DQ368381 (pGRW6); KT282462 (pERIRUB01) [23, 120] P. hermani Not available Not available P. huffi Not available Not available P. polymorphum Not available Not available Only DNA sequences, for which parasite species identity was supported by morphological analysis are included in this table References of articles containing discussion of molecular characterization and morphological features of parasite species same lineage after 3–4 passages via the infected blood cuckoo (Fig. 6a) and after the first passage in the Eura - inoculation in passeriform birds (G. Valkiūnas, unpub- sian siskin Carduelis spinus (Passeriformes) (Fig. 6b) lished). Molecular testing showed that the parasite line- and several subsequent passages in siskins (Fig. 6c, d) age was the same. Pictures of erythrocytic meronts of illustrate this change. These experimental data indicate the same isolate of the lineage pGRW2 in the Common that malaria parasites which do not possess globules in Valkiūnas and Iezhova Malar J (2018) 17:212 Page 15 of 24 Table 8 List of synonyms of Plasmodium species of birds Table 8 (continued) parasites were described in co-infection with Haemamoeba parasites, including Synonymous name and references of original Valid name P. relictum, and description of blood stages were fragmentary [56, 57]. Blood description stages of all these parasites do not have unique characters, which could help to distinguish them from P. relictum. Plasmodium bioccai, P. coluzzii, P. dorsti, P. Plasmodium alaudae  P. relictum (partim) ginsburgi, P. relictum quentini are considered as synonyms of P. relictum P. alloelongatum  P. elongatum d Synonymous status of P. mohammedi was specified in Table 9 (see the footnote “f ”) P. bioccai  P. relictum P. biziurae  P. relictum P. capistrani  P. relictum Table 9 List of species names of bird malaria parasites P. centropi  P. cathemerium (partim) belonging to the categories of nomen nudum, nomen P. chloropsidis  P. relictum dubium, species inquirenda and incertae sedis P. coluzzii  P. relictum Name and references Status P. dorsti  P. relictum c b P. ginsburgi  P. relictum Plasmodium alaudae [57, 126] Species inquirenda P. arachnidi  Species inquirenda P. heroni  P. circumflexum P. bambusicolai  Species inquirenda P. huffi  P. nucleophilum P. beaucournui  Species inquirenda P. japonicum  P. juxtanucleare P. bigueti  Species inquirenda P. metastaticum  P. gallinaceum P. buteonis  Species inquirenda P. grassii  P. relictum P. coggeshalli  Species inquirenda P. inconstans  P. relictum P. conturnixae  Nomen nudum P. maior  P. relictum P. corradettii  Nomen dubium P. merulae  P. vaughani d P. danilewskyi [152, 153] Incertae sedis P. mohammedi  P. rouxi Plasmodium dherteae  Species inquirenda P. muniae  P. relictum P. gallinulae  Incertae sedis P. majoris  P. relictum (partim) P. gambeli  Nomen nudum P. oti  P. hexamerium P. ghadiriani  Species inquirenda P. passeris  P. relictum P. golvani  Species inquirenda P. paddae  P. relictum P. herodiadis  Species inquirenda P. pericrocoti  P. relictum P. holti  Nomen nudum P. ploceii  P. relictum P. jiangi  Species inquirenda P. relictum quentini  P. relictum P. jeanriouxi  Species inquirenda P. spheniscidae  P. relictum P. lagopi  Species inquirenda P. tumbayaensis  P. vaughani P. lairdi  Nomen nudum P. tenuis  P. vaughani P. lenoblei  Species inquirenda Plasmodium wasielewskii  P. subpraecox P. malariae raupachi  Incertae sedis Plasmodium species synonymous names published before 2000 were justified P. manwelli  Nomen nudum in  d P. ninoxi  Species inquirenda According to the original description , P. alloelongatum is similar to P. P. noctuae [3, 126] Species inquirenda elongatum, but differs from the latter species mainly due to two characters: P. pachysomum  Species inquirenda (1) the erythrocytic meront progeny is limited to 6 (predominantly 6–12 in P. elongatum), and (2) the undulating or rugged outlines and tapering gametocyte P. papernai  Species inquirenda ends, which might extend into a distal spine or filaments. The irregularity of e Plasmodium pfefferi  Species inquirenda gametocyte shape (Fig. 4p) and presence of ameboid outgrowth (Fig. 4q) has P. praecox  Nomen nudum been reported and illustrated in P. elongatum (the closely related lineages pGRW6 and pERIRUB01), but have rarely pointed out in descriptions of this P. reniai  Species inquirenda parasite [1, 23, 120]. Furthermore, the ameboid outgrowths in gametocytes P. rousseloti  Species inquirenda were seen in the neohapantotype of P. elongatum (blood slide no. 216, the P. rouxi, as published in  Species inquirenda (probably a new Plasmo- Natural History Museum, London). The number of nuclei in mature erythrocytic dium species) meronts is variable in P. elongatum during development in different host cells and avian hosts, and it is often ≤ 6 [1, 23, 120]. Plasmodium elongatum has been P. sergentorum  Species inquirenda characterized molecularly (Table 7), and it has been reported in numerous b P. snounoui  Species inquirenda bird species belonging to different orders both by microscopic examination P. spartani  Nomen nudum of blood films and PCR-based testing, including species of Accipitriformes and Falconiformes . Based on available information, Plasmodium alloelongatum P. stellatum  Species inquirenda cannot be distinguished is considered as a synonym of P. elongatum P. struthionis  Incertae sedis Observation of blood stages in various bird species experimentally infected b P. tranieri  Species inquirenda with single infections of Plasmodium relictum lineages pSGS1 and pGRW11, P. venkataramiahii  Nomen nudum which are closely related and widespread in Europe, show that main reported P. bioccai, P. coluzzii, P. dorsti, P. ginsburgi, P. relictum quentini blood stages (meronts Nomenclature status of the species names published before 2000 was justified and gametocytes) are present in these parasite lineages [67, 71, 117]. These in  Valkiūnas and Iezhova Malar J (2018) 17:212 Page 16 of 24 Table 9 (continued) Table 9 (continued) Plasmodium beaucournui, P. bigueti, P. coggeshalli, P. dherteae, P. ghadiriani, P. golvani, possess refractive globules (Fig. 3f, j) and gametocytes possess few large P. jeanriouxi, P. lenoblei, P. papernai, P. reniai, P. snounoui, P. tranieri were named and (Fig. 4o) pigment granules. The latter character is an important feature of P. described, and P. alaudae was re-described from individual birds co-infected with rouxi. Based on available information, the parasite described in  as ‘P. rouxi’ parasites belonging to subgenera Haemamoeba, Giovannolaia and Novyella [56, cannot be attributed to P. rouxi and is considered as a species inquirenda. The 57, 65, 149]. The authors of the original descriptions have grouped the blood stages parasite described by Paperna et al.  is characterized by presence of (1) the visible in blood films and attributed them to different species provisionally, which relatively prominent cytoplasm in growing meronts and (2) tiny size of pigment is particularly obvious in case of parasites with elongate gametocytes. This makes granules in gametocytes, so might belong to a new Plasmodium species. species description and validation of parasite names questionable. Only single cells Additional investigation is needed to answer this question. In the same study, (erythrocytic meronts) were selected as holotypes in these parasite descriptions. Paperna et al.  described a new species Plasmodium mohammedi, which However, due to morphological variation of blood stages of Plasmodium and presence was reported, Passer domesticus (the common host of P. rouxi in Mediterranian of parasites at different stages of growth in each blood film, such methodology of region ). Blood stages of P. mohammedi are indistinguishable from P. rouxi designation of the type material can work only in case of exceptionally distinctive , particularly due to the presence of refractive globules in erythrocytic cell characters, which is not the case in all these parasite descriptions, particularly meronts and large pigment granules in gametocytes (see Figs. 18–21 in ). belonging to subgenus Haemamoeba. Molecular characterization of all these parasites Plasmodium mohammedi is a synonym of P. rouxi. Molecular identification of P. is unavailable. It is clear from the original descriptions, that many individual birds rouxi (lineage pPADOM16) was developed . Application of the barcoding were infected by representatives of several subgenera. However, the reported blood indicates that the details of disposition of nuclei in erythrocytic meronts during stages were selected and attributed to certain species without providing convincing different infections, particularly in different avian hosts, is variable in P.rouxi, explanations, making identifications difficult or even impossible based on available but binuclear “bow-tie” form parasites often are present (Fig. 3f ) and can be information. Co-infections of Plasmodium parasites belonging to different subgenera used for this parasite species identification. Additionally, presence of few large are common in wildlife, and the described cases of co-infections with several malaria pigment granules in mature gametocytes also is a characteristic feature, and parasites are not unpredictable . However, description of new species from such it recommended to use for distinguishing P. rouxi infection (Fig. 4o) from other co-infections hardly possible if the unique morphological characters of blood stages Novyella parasites producing tetranuclear erythrocytic meronts are absent, which is the case with P. beaucournui, P. bigueti, P. coggeshalli, P. dherteae, P. ghadiriani, P. golvani, P. jeanriouxi, P. lenoblei, P. papernai, P. reniai, P. snounoui, P. tranieri and also in re-description of P. alaudae. These parasites are considered as species inquirenda. Recent molecular studies provided molecular markers for distinguishing natural hosts might develop this structure after artifi - blood stages of Plasmodium species (Table 7). Examination of blood films from cial passages via infected blood inoculation in unusual experimental infections shows variations in morphological characters of same parasite lineages in different avian hosts, calling for careful application of minor differences avian hosts. In other words, this feature hardly can be in blood stage morphology in avian malaria parasite taxonomy, particularly during used in taxonomy of avian Plasmodium parasites at sub- co-infections genus level. It is preferable to limit use of the feature of Based on available information , P. buteonis cannot be distinguished from P. circumflexum and other similar parasites of Giovannolaia ( Plasmodium gabaldoni, absence or presence of globules in erythrocytic meronts Plasmodium homocircumflexum). The main feature, which has been noted to to identification of natural infections at species level, on distinguish P. buteonis from P. circumflexum in the original description , is the which the taxonomic validity of this feature also needs presence up to 36 nuclei in mature erythrocytic meronts of the former. Plasmodium circumflexum produce less number of nuclei in mature meronts. However, the to be tested. Experimental sporozoite-induced infections description of P. buteonis is based on high parasitemia (6.6%), with numerous of same parasites lineages possessing and not possess- multiple infections of the same erythrocytes, so it is difficult to rule out that 2 mature meronts were present in same cell in case of so great number of merozoites. ing globules in different avian hosts might help to answer Additionally, parasite morphology often changes during high parasitemia, so such the question about taxonomic value of this feature. Until samples should be carefully used in taxonomical descriptions. Plasmodium buteonis additional information is available, Papernaia is consid- might be a valid name, but more research is needed to prove its validity. Molecular characterization of this parasite is absent, but is essential to solve the question about ered as a synonym of subgenus Novyella. its validity Plasmodium ninoxi was described from owl Ninox scutulata in co-infection Conclusion with Haemoproteus sp. . Only one erythrocyte with 2 binuclear growing meronts was detected; no other data about merogony in the blood were Based on available morphological data and DNA provided. Plasmodium ninoxi gametocytes were reported to be rounded. Based sequence information, 55 species of avian Plasmodium on available information, it seems that infected blood was exposed to air, which stimulated rounding-up of haemoproteid gametocytes , which were parasites can be readily distinguished. Species of subge- attributed to P. ninoxi. DNA sequence was provided (AY099035.1), and it belongs nus Novyella predominate among them. Dichotomous to Plasmodium sp. Plasmodium ninoxi description is incomplete. Re-description keys for identification of these parasites were compiled is needed, and it is possible due to available sequence information. The most similar cytb sequence belong to P. gallinaceum, P. relictum and P. circumflexum allowing identification of these pathogens using mor - Descriptions of P. pachysomum, P. pfefferi, P. sergentorum, P. stellatum  are phological features of their blood stages. The major - incomplete. Information about morphology of gametocytes is absent. Molecular ity of described avian Plasmodium species are mainly characterization is unavailable. Species identification is questionable based on the available information transmitted in countries with warm climates. The Paperna et al.  published re-description of P. rouxi from non-type avian obstacles for their global spread remain insufficiently host (Alauda arvensis, Alaudidae instead of Passer hispaniolensis, Passeridae understood, mainly because of limited information on whose is the type host). The re-description is based on samples, which were collected beyond of the type locality (France, instead of Algeria which is the life cycles and vectors of the majority of described par- type locality). This contradicts the Article 75.3.6 of the International Code of asites of tropical birds. The lists of synonymous names Zoological Nomenclature . Additionally, according to , the erythrocytic as well as names of the categories species inquirenda meronts of the parasite from A. arvensis do not possess refractive globules and gametocytes possess few tiny pigment granules (Figs. 8, 9 in ). These and incertae sedis should be considered in future taxo- are not characters of P. rouxi, which was described by Sergent et al. . nomic work of avian malaria parasite at species level. Sergent’s original material from Algiers labelled “2198, 26.4.28, Institut Pasteur d’Algérie” is available in the Natural History Museum, London. Examination of The majority of described Plasmodium parasites have this blood film showed that numerous erythrocytic meronts of this parasite not been characterized using molecular markers, which Valkiūnas and Iezhova Malar J (2018) 17:212 Page 17 of 24 Fig. 2 Morphological features of erythrocytic meronts and their host cells of avian Plasmodium parasites, which are used for Haemamoeba, Giovannolaia and Huffia species identification. Growing (a–c, f–h, l–p) and mature (d, e, i–k) meronts at different stages of their development. Note presence of the plentiful cytoplasm and large nuclei in early growing meronts (a, b, f–h, m–p), marked vacuolization of the cytoplasm (f–h), elongate shape of mature merozoites (k), presence of meronts in erythroblasts (i, l–n) and other immature red blood cells (k, o, p), and distinct smooth outline in growing erythrocytic meronts (m, n). Short simple arrows—vacuoles. Wide triangle arrowheads—the cytoplasm. Other symbols are as in Fig. 1. Explanations are given in the text Valkiūnas and Iezhova Malar J (2018) 17:212 Page 18 of 24 Fig. 3 Morphological features of erythrocytic meronts and their host cells of avian Plasmodium parasites, which are used for Novyella and Giovannolaia species identification. Trophozoites (a–d) and erythrocytic meronts (e–y) on different stages of maturation. Note presence of large vacuoles (a, e, m), refractive small globules (f, h–j), bluish non-refractive globules (b, k, l), fan-like mature meronts (o, v), strictly nucleopilic position (n, t), the scanty (nearly invisible) cytoplasm (a, b, e–l) and the prominent (readily visible) cytoplasm (d, x) in parasites on different stages of their development. Triangle wide long arrows—refractive globules. Triangle wide short arrows—bluish (non-refractive) globules. Other symbols are as in Fig. 1. Explanations are given in the text Valkiūnas and Iezhova Malar J (2018) 17:212 Page 19 of 24 Fig. 4 Morphological features of gametocytes and their host cells of avian Plasmodium parasites, which are used for species identification. Macrogametocytes (a–g, k–u, w–y) and microgametocytes (h–j, v). Note long outgrowth (f), terminal position of pigment granules (e) and nucleus (g), granular (l, m) and vacuolated (n) appearance of the cytoplasm, slender (p–r) and circumnuclear (s) shapes of gametocytes, clumps of pigment granules located near the parasite margin (t, w), distinct smooth outline of nucleus (y). Symbols as in Figs. 1, 2, 3. Explanations are given in the text Valkiūnas and Iezhova Malar J (2018) 17:212 Page 20 of 24 Fig. 5 Morphological features of blood stages and their host cells of avian Plasmodium parasites, which are used for species identification. Young trophozoite (a) and gametocyte (b), growing erythrocytic meronts (c, d, j, u), mature erythrocytic meronts (f, p–s, w), and mature gametocytes (e, g–i, k–o, t, v, x, y). Note presence of long outgrowths (a–c), terminal position of nuclei in meront (d), slender shape of gametocyte (e), aggregation of pigment granules at one end of gametocyte (f), rod-like pigment granules (n), large vacuoles (g, j, u), refractive globules in gametocyte (h), oblique position of gametocytes in erythrocytes (i, o), strictly nucleophilic erythrocytic meronts (q), residual cytoplasm in erythrocytic meronts (r, s), rounded shape of infected erythrocytes (p, w–y). Triangle long arrows—residual body in mature meront. Symbols as in Figs. 1, 2, 3, 4. Explanations are given in the text Valkiūnas and Iezhova Malar J (2018) 17:212 Page 21 of 24 Fig. 6 Maturing erythrocytic meronts of Plasmodium ashfordi (lineage pGRW2) in naturally infected the Common cuckoo Cuculus canorus (a) and experimentally infected Eurasia siskin Carduelis spinus (b–d) during the first (b) and 3–4th (c, d) passages of infected blood. Note that refractive globules were absent in erythrocytic meronts during the natural infection (a) and the first passage of the experimental infection (b), but develop in subsequent passages of the same strain in Eurasian siskin. Symbols are as in Figs. 1 and 3 3. Bennett GF, Whiteway M, Woodworth-Lynas C. A host-parasite cata- development is an essential task for current avian logue of the avian haematozoa. Occasional Papers in Biology. St. John’s: malaria researchers. Memorial University of Newfoundland; 1982. p. 243. 4. Atkinson CT, Thomas NJ, Hunter DB. Parasitic diseases of wild birds. Authors’ contributions Oxford: Wiley-Blackwell; 2008. GV collected published articles and collection material, analysed the literature 5. Clark NJ, Clegg SM, Lima MR. A review of global diversity in avian hae- data and wrote the manuscript; GV and TAI analysed preparations of the blood mosporidians (Plasmodium and Haemoproteus: Haemosporida): new stages; TAI and GV prepared plates of images. Both authors read and approved insights from molecular data. Int J Parasitol. 2014;44:329–38. the final manuscript. 6. Hellgren O, Atkinson CT, Bensch S, Albayrak T, Dimitrov D, Ewen JG, et al. Global phylogeography of the avian malaria pathogen Plasmodium relictum based on MSP1 allelic diversity. Ecography. 2015;38:842–50. Acknowledgements 7. Sehgal RN. Manifold habitat effects on the prevalence and diversity of This article benefited from comments made by Richard W. Ashford and avian blood parasites. Int J Parasitol Parasites Wildl. 2015;4:421–30. Carolina R. F. Chagas. We thank R. Adlard, E. Hoberg, A. Warren, I. Landau, C. 8. Valkiūnas G. Avian malaria parasites and other Haemosporidia. Boca Atkinson and N.E. Matta for assistance in accessing parasite material and D. Raton: CRC; 2005. Bukauskaitė, M. Ilgūnas, V. Palinauskas and R. Žiegytė, for participation in field 9. Santiago-Alarcon D, Palinauskas V, Schaefer HM. Diptera vectors of work during collection of samples. avian Haemosporidian parasites: untangling parasite life cycles and their taxonomy. Biol Rev Camb Philos Soc. 2012;87:928–64. Competing interests 10. Njabo K, Cornel AJ, Sehgal RNM, Loiseau C, Buermann W, Harrigan RJ, The authors declare that they have no competing interests. et al. Coquillettidia (Culicidae, Diptera) mosquitoes are natural vectors of avian malaria in Africa. Malar J. 2009;8:193. Availability of data and materials 11. Ejiri H, Sato Y, Kim KS, Tsuda Y, Murata K, Saito K, et al. Blood meal All data generated during this study are included in this published article. identification and prevalence of avian malaria parasite in mosquitoes collected at Kushiro Wetland, a subarctic zone of Japan. J Med Entomol. Consent for publication 2011;48:904–8. Not applicable. 12. Garnham PCC. Malaria in its various vertebrate hosts. In: Kreier JP, editor. Malaria. Part 1. Epidemiology, chemotherapy, morphology and Ethics approval and consent to participate metabolism. New York: Academic Press; 1980. p. 95–144. Not applicable. 13. Sherman IW. Malaria: parasite biology, pathogenesis, and protection. Washington: ASM; 1998. Funding 14. Cowman AF, Healer J, Marapana D, Marsh K. Malaria: biology and This study was funded by the Research Council of Lithuania (No. disease. Cell. 2016;167:610–24. MIP-045/2015). 15. Valkiūnas G, Žiegytė R, Palinauskas V, Bernotienė R, Bukauskaitė D, Ilgūnas M, et al. Complete sporogony of Plasmodium relictum (lineage pGRW4) in mosquitoes Culex pipiens pipiens, with implications on avian Publisher’s Note malaria epidemiology. Parasitol Res. 2015;144:3075–85. Springer Nature remains neutral with regard to jurisdictional claims in pub- 16. Žiegytė R, Bernotienė R, Bukauskaitė D, Palinauskas V, Iezhova TA, lished maps and institutional affiliations. Valkiūnas G. Complete sporogony of Plasmodium relictum (line- ages pSGS1 and pGRW11) in mosquito Culex pipiens pipiens form Received: 24 March 2018 Accepted: 15 May 2018 molestus, with implications to avian malaria epidemiology. J Parasitol. 2014;100:878–82. 17. 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. References 18. Loiseau C, Harrigan RJ, Cornel AJ, Guers SL, Dodge M, Marzec T, et al. 1. Garnham PCC. Malaria parasites and other Haemosporidia. Oxford: First evidence and predictions of Plasmodium transmission in Alaskan Blackwell; 1966. bird populations. PLoS ONE. 2012;7:e44729. 2. Seed TM, Manwell RD. Plasmodia of birds. In: Kreier JP, editor. Parasitic 19. Schoener ER, Banda M, Howe L, Castro IC, Alley MR. Avian malaria in protozoa, vol III. Gregarines, Haemogregarines, Coccidia, Plasmodia, and New Zealand. NZ Vet J. 2014;62:189–98. Haemoproteids. New York: Academic Press; 1977. p. 311–57. 20. Marzal A. Recent advances in studies on avian malaria parasites. In: Okwa OO, editor. Malaria parasites. In Tech: Rijeka; 2012. p. 135–58. Valkiūnas and Iezhova Malar J (2018) 17:212 Page 22 of 24 21. Bensch S, Hellgren O, Pérez-Tris J. MalAvi: a public database of malaria 41. Vanstreels RE, da Silva-Filho RP, Kolesnikovas CK, Bhering RC, Ruoppolo parasites and related haemosporidians in avian hosts based on mito- V, Epiphanio S, et al. Epidemiology and pathology of avian malaria in chondrial cytochrome b lineages. Mol Ecol Resour. 2009;9:1353–8. penguins undergoing rehabilitation in Brazil. Vet Res. 2015;46:30. 22. Zehtindjiev P, Križanauskienė A, Scebba S, Dimitrov D, Valkiūnas G, 42. Valkiūnas G, Ashford RW. Natural host range is not a valid taxonomic Heggemann A, et al. Haemosporidian infections in skylarks (Alauda character. Trends Parasitol. 2002;18:528–9. arvensis): a comparative PCR-based and microscopy study on the 43. Kim KS, Tsuda Y, Yamada A. Blood meal identification and detection of parasite diversity and prevalence in southern Italy and the Netherlands. avian malaria parasite from mosquitoes (Diptera: Culicidae) inhabiting Eur J Wildl Res. 2012;58:335–44. coastal areas of Tokyo Bay, Japan. J Med Entomol. 2009;46:1230–4. 23. Palinauskas V, Žiegytė R, Iezhova TA, Ilgūnas M, Bernotienė R, Valkiūnas 44. Dinhopl N, Nedorost N, Mostegl MM, Weissenbacher-Lang C, Weis- G. Description, molecular characterisation, diagnostics and life cycle of senböck H. In situ hybridization and sequence analysis reveal an Plasmodium elongatum (lineage pERIRUB01), the virulent avian malaria association of Plasmodium spp. with mortalities in wild passerine birds parasite. Int J Parasitol. 2016;46:697–707. in Austria. Parasitol Res. 2015;114:1455–62. 24. Ilgūnas M, Bukauskaitė D, Palinauskas V, Iezhova TA, Dinhopl N, 45. Valkiūnas G, Iezhova TA, Shapoval AP. High prevalence of blood parasites Nedorost N, et al. Mortality and pathology in birds due to Plasmo- in hawfinch Coccothraustes coccothraustes. J Nat Hist. 2003;37:2647–52. dium (Giovannolaia) homocircumflexum infection, with emphasis on 46. Valkiūnas G, Bensch S, Iezhova TA, Križanauskienė A, Hellgren O, Bolsha- the exoerythrocytic development of avian malaria parasites. Malar J. kov CV. Nested cytochrome b polymerase chain reaction diagnostics 2016;15:256. underestimate mixed infections of avian blood haemosporidian 25. Valkiūnas G, Iezhova TA. Exo-erythrocytic development of avian malaria parasites: microscopy is still essential. J Parasitol. 2006;92:418–22. and related haemosporidian parasites. Malar J. 2017;16:101. 47. Martínez J, Martínez-De La Puente J, Herrero J, Del Cerro S, Lobato 26. Corradetti A, Garnham PCC, Neri L, Scanga M, Cavallini C. A redescrip- E, Rivero-De Aguilar J, Cerro S, Lobato E, Rivero-De Aguilar J, et al. A tion of Plasmodium (Haemamoeba) relictum (Grassi and Feletti, 1891). restriction site to differentiate Plasmodium and Haemoproteus infec- Parassitologia. 1970;12:1–10. tions in birds: on the inefficiency of general primers for detection of 27. Bennett GF, Warren M, Cheong WH. Biology of the Malaysian strain of mixed infections. Parasitology. 2009;136:713–22. Plasmodium juxtanucleare Versiani and Gomes, 1941. II. The sporogonic 48. Bernotienė R, Palinauskas V, Iezhova TA, Murauskaitė D, Valkiūnas G. stages in Culex (Culex) sitiens Wiedmann. J Parasitol. 1966;52:647–52. Avian haemosporidian parasites (Haemosporida): a comparative analy- 28. Martinsen ES, Perkins SL, Schall JJ. A three-genome phylogeny of sis of different polymerase chain reaction assays in detection of mixed malaria parasites (Plasmodium and closely related genera): evolu- infections. Exp Parasitol. 2016;163:31–7. tion of life-history traits and host switched. Mol Phylogenet Evol. 49. Mantilla JS, González AD, Valkiūnas G, Moncada LI, Matta NE. Descrip- 2008;47:261–73. tion and molecular characterization of Plasmodium (Novyella) unalis sp. 29. Outlaw DC, Ricklefs RE. Species limits in avian malaria parasites (Hae- nov. from the Great Thrush (Turdus fuscater) in highland of Colombia. mosporida): how to move forward in the molecular era. Parasitology. Parasitol Res. 2013;112:4193–204. 2014;141:1223–32. 50. Silveira P, Belo NO, Lacorte GA, Kolesnikovas CKM, Vanstreels RET, 30. Perkins SL. Malaria’s many mates: past, present and future of the sys- Steindel M, et al. Parasitological and new molecular-phylogenetic tematics of the order Haemosporida. J Parasitol. 2014;100:11–25. characterization of the malaria parasite Plasmodium tejerai in South 31. Outlaw RK, Counterman B, Outlaw DC. Differential patterns of molecu- American penguins. Parasitol Int. 2013;62:165–71. lar evolution among Haemosporidian parasite groups. Parasitology. 51. Savage AF, Ariey F, Greiner EC. A new species of Plasmodium from 2015;142:612–22. Malagasy vangas. J Parasitol. 2005;9:926–30. 32. Bensch S, Canbäck B, DeBarry JD, Johansson T, Hellgren O, Kissinger JC, 52. Valkiūnas G, Iezhova TA, Loiseau C, Chasar A, Smith TB, Sehgal RNM. New spe- et al. The genome of Haemoproteus tartakovskyi and its relationship to cies of haemosporidian parasites (Haemosporida) from African rainforest human malaria parasites. Genome Biol Evol. 2016;8:1361–73. birds, with remarks on their classification. Parasitol Res. 2008;103:1213–28. 33. Pacheco AM, Matta NE, Valkiūnas G, Parker PG, Mello B, Stanley CE 53. Paperna I, Yosef R, Landau I. Plasmodium spp. in raptors on the Eurasian- Jr, et al. Mode and rate of evolution of haemosporidian mitochon- African migration route. Parasite. 2007;14:313–22. drial genomes: timing the radiation of avian parasites. Mol Biol Evol. 54. Paperna I, Yosef R, Chavatte JM, Grill H, Landau I. Species of Plasmodium 2018;35:383–403. of passerine birds with four nuclei, with description of new species. 34. Adams Y, Kuhnrae P, Higgins MK, Ghumra A, Rowe JA. Rosetting Acta Parasitol. 2008;2008(53):227–36. Plasmodium falciparum-infected erythrocytes bind to human brain 55. Valkiūnas G, Iezhova TA, Loiseau C, Smith TB, Sehgal RNM. New malaria microvascular endothelial cells in vitro, demonstrating a dual adhesion parasites of the subgenus Novyella in African rainforest birds, with phenotype mediated by distinct P. falciparum erythrocyte membrane remarks on their high prevalence, classification and diagnostics. Parasi- protein 1 domains. Infect Immun. 2014;82:949–59. tol Res. 2009;104:1061–77. 35. Palinauskas V, Žiegytė R, Ilgūnas M, Iezhova TA, Bernotienė R, Bolshakov 56. Chavatte JM, Chiron F, Chabaud A, Landau I. Probable speciations by C, et al. Description of the first cryptic avian malaria parasite, Plasmo - “host-vector ‘fidelity’”: 14 species of Plasmodium from magpies. Parasite. dium homocircumflexum n. sp., with experimental data on its virulence 2007;14:21–37 (in French). and development in avian hosts and mosquitoes. Int J Parasitol. 57. Chavatte JM, Grés V, Snounou G, Chabaud A, Landau I. Plasmo- 2015;45:51–62. dium (Apicomplexa) of the skylark (Alauda arvensis). Zoosystema. 36. Huchzermeyer FW, van der Vyver FH. Isolation of Plasmodium circum- 2009;31:369–83. flexum from wild guineafowl (Numida meleagris) and the experimental 58. Walther EL, Valkiūnas G, González AD, Matta NE, Ricklefs RE, Cornel A, infection in domestic poultry. Avian Pathol. 1991;20:213–23. et al. Description, molecular characterization, and patterns of distribu- 37. Huchzermeyer FW. Pathogenicity and chemotherapy of Plasmodium tion of a widespread New World avian malaria parasite (Haemosporida: durae in experimentally infected domestic turkeys. Onderstepoort J Vet Plasmodiidae), Plasmodium (Novyella) homopolare sp. nov. Parasitol Res. Res. 1993;60:103–10. 2014;113:3319–32. 38. Palinauskas V, Valkiūnas G, Bolshakov CV, Bensch S. Plasmodium relictum 59. Zehtindjiev P, Križanauskienė A, Bensch S, Palinauskas V, Asghar M, Dim- (lineage P-SGS1): effects on experimentally infected passerine birds. Exp itrov D, et al. A new morphologically distinct avian malaria parasite that Parasitol. 2008;120:372–80. fails detection by established PCR-based protocols for amplification of 39. Vanstreels RE, Kolesnikovas CK, Sandri S, Silveira P, Belo NO, Ferreira the cytochrome b gene. J Parasitol. 2012;98:657–65. Junior FC, et al. Outbreak of avian malaria associated to multiple species 60. Telford SR. The hemoparasites of the reptilian. Boca Raton: CRC; 2009. of Plasmodium in magellanic penguins undergoing rehabilitation in 61. Lainson R. Atlas of protozoan parasites of the Amazonian fauna of Brazil. southern Brazil. PLoS ONE. 2014;9:e94994. Haemosporida of reptiles, vol. 1. Ananindeua: Instituto Evandro Chagas; 40. Dimitrov D, Palinauskas V, Iezhova TA, Bernotienė R, Ilgūnas M, 2012. Bukauskaitė D, et al. Plasmodium spp.: an experimental study on verte- 62. Gabaldon A, Ulloa G, Zerpa N. Fallisia (Plasmodioides) neotropicalis brate host susceptibility to avian malaria. Exp Parasitol. 2015;148:1–16. subgen. nov. sp. nov. from Venezuela. Parasitology. 1985;90:217–25. Valkiūnas and Iezhova Malar J (2018) 17:212 Page 23 of 24 63. Campbell TW. Avian hematology and cytology. Ames: Iowa State 85. Grassi B, Feletti R. Malariaparasiten in den Vögeln. Centralbl Bakteriol University Press; 1995. Parasitenkd. 1891;9:403–9, 429–33, 461–7. 64. Valkiūnas G, Iezhova TA, Križanauskienė A, Palinauskas V, Sehgal RNM, 86. Hartman E. Three species of bird malaria, Plasmodium praecox, P. cath- Bensch S. A comparative analysis of microscopy and PCR-based detec- emerium n. sp. and P. inconstans n. sp. Arch Protistenkd. 1927;60:1–7. tion methods for blood parasites. J Parasitol. 2008;94:1395–401. 87. Corradetti A, Verolini F, Neri I. Plasmodium (Haemamoeba) giovannolai n. 65. Landau I, Chabaud AG, Bertani S, Snounou G. Taxonomic status and re- sp. parassita di Turdus merula. Parassitologia. 1963;5:11–8. description of Plasmodium relictum (Grassi et Feletti, 1891), Plasmodium 88. Huff CG. A new variety of Plasmodium relictum from the robin. J Parasi- maior Raffaele, 1931, and description of P. bigueti n. sp. in sparrows. tol. 1937;23:400–4. Parassitologia. 2003;45:119–23. 89. Gabaldon A, Ulloa G. Plasmodium (Haemamoeba) tejerai sp. n. del pavo 66. Valkiūnas G, Ilgūnas M, Bukauskaitė D, Palinauskas V, Bernotienė R, domésstico (Meleagris gallopavo) de Venezuela. Bol Dir Malariol San Iezhova TA. Molecular characterization and distribution of Plasmo- Amb. 1977;17:255–73. dium matutinum, a common avian malaria parasite. Parasitology. 90. Lucena D. Malária aviária I—Plasmodium lutzi n. sp. Parasita da Saracura 2017;144:1726–35. (Aramides cajanea cajanea, Müller). Bull Biol (São Paulo). 1939;4:27–31. 67. Palinauskas V, Kosarev V, Shapoval A, Bensch S, Valkiūnas G. Comparison 91. Mantilla JS, Matta NE, Pacheco MA, Escalante AA, Gonzalez AD, of mitochondrial cytochrome b lineages and morphospecies of two Moncada LI. Identification of Plasmodium (Haemamoeba) lutzi (Lucena, avian malaria parasites of the subgenera Haemamoeba and Giovan- 1939) from Turdus fuscater (Great Thrush) in Colombia. J Parasitol. nolaia (Haemosporida: Plasmodiidae). Zootaxa. 2007;1626:39–50. 2013;99:662–8. 68. Marzal A, Ricklefs RE, Valkiūnas G, Albayrak T, Arriero E, Bonneaud C, 92. Schwetz J. Sur un Plasmodium aviaire à formes de division allongées, et al. Diversity, loss and gain of malariae parasites in a globally invasive Plasmodium fallax, n. sp. Arch Inst Pasteur Alger. 1930;8:289–96. bird. PLoS ONE. 2011;6:e21905. 93. Manwell RD, Kuntz RE. Plasmodium hegneri n. sp. from the European 69. Beadell JS, Ishtiaq F, Covas R, Melo M, Waren BH, Atkinson CT, et al. Teal Anas c. crecca in Taiwan. J Protozool. 1966;13:437–40. Global phylogeographic limits of Hawaii’s avian malaria. Proc R Soc 94. Huang J-C. A new species of the genus Plasmodium—Plasmodium Lond B Biol Sci. 2006;273:2935–44. leanucleus (Eucoccidia: Plasmodiidae). Acta Vet Zoot Sinica. 70. Bueno MG, Lopez RP, de Menezes RM, de Costa-Nascimento MJ, Lima 1991;16:257–62. GF, Araújo RA, et al. Identification of Plasmodium relictum causing mor - 95. Bray RS. On the parasitic Protozoa of Liberia. VII—Haemosporidia of tality in penguins (Spheniscus magellanicus) from São Paulo Zoo, Brazil. owls. Arch Inst Pasteur Alger. 1962;40:201–7. Vet Parasitol. 2010;173:123–7. 96. Manwell RD. Plasmodium octamerium n. sp., an avian malaria 71. Valkiūnas G, Zehtindjiev P, Hellgren O, Ilieva M, Iezhova TA, Bensch S. parasite from the pintail whydah bird Vidua macroura. J Protozool. Linkage between mitochondrial cytochrome b lineages and morphos- 1968;15:680–5. pecies of two avian malaria parasites, with a description of Plasmodium 97. Kikuth W. Immunbiologische und chemotherapeutische Studien an (Novyella) ashfordi sp. nov. Parasitol Res. 2007;100:1311–22. verschiedenen Stämmen von Vogelmalaria. Zentralbl Bakteriol Para- 72. Valkiūnas G, Ilgūnas M, Bukauskaitė D, Žiegytė R, Bernotienė R, Jusys sitenkd Infektionskr Hyg I Abt Orig. 1931;121:401–9. V, et al. Plasmodium delichoni n. sp.: description, molecular characteri- 98. Coggeshall LT. Plasmodium lophurae, a new species of malaria parasite sation and remarks on the exoerythrocytic merogony, persistence, pathogenic for the domestic fowl. Am J Hyg. 1938;27:615–8. vectors and transmission. Parasitol Res. 2016;115:2625–36. 99. Garnham PCC. A new malaria parasite of pigeons and ducks from 73. Ricklefs RE, Soares L, Ellis VA, Latta SC. Avian migration and the distribu- Venezuela. Protistologica. 1977;13:113–25. tion of malaria parasites in New World passerine birds. J Biogeogr. 100. Manwell RD. How many species of avian malaria parasites are there? J 2017;44:1113–23. Parasitol. 1934;20:334. 74. Zehtindjiev P, Ilieva M, Westerdahl H, Hansson B, Valkiūnas G, Bensch 101. Muniz J, de Soares RRL. Nota sôbre um parasita do gênero Plasmodium S. Dynamics of parasitemia of malaria parasites in a naturally and encontrado no Ramphastos toco Müller, 1776, “Tugano-Açu”, e diferente experimentally infected migratory songbird, the great reed warbler do Plasmodium huffi: Plasmodium pinottii n. sp. Rev Bras Malariol. Acrocephalus arundinaceus. Exp Parasitol. 2008;119:99–110. 1954;6:611–7. 75. Palinauskas V, Valkiūnas G, Bolshakov CV, Bensch S. Plasmodium relictum 102. Guindy E, Hoogstraal H, Mohammed AHH. Plasmodium garnhami sp. (lineage SGS1) and Plasmodium ashfordi (lineage GRW2): the effects nov. from the Egyptian hoopoe (Upupa epops major Brehm). Trans R Soc of the co-infection on experimentally infected passerine birds. Exp Trop Med Hyg. 1965;59:280–4. Parasitol. 2011;127:527–33. 103. Manwell RD. A new species of avian Plasmodium. J Protozool. 76. Escalante AA, Freeland DE, Collins WE, Lal AA. The evolution of primate 1962;9:401–3. malaria parasites based on the gene encoding cytochrome b from the 104. Herman CM. Plasmodium durae, a new species of malaria parasite from linear mitochondrial genome. Proc Natl Acad Sci USA. 1998;95:8124–9. the common turkey. Am J Hyg. 1941;34:22–6. 77. Bensch S, Stjernman M, Hasselquist D, Östman Ö, Hansson B, Wester- 105. Shillinger JE. Diseases of wildlife and their relationship to domestic dahl H, et al. Host specificity in avian blood parasites: a study of Plasmo - livestock. Washington: USDA Yearbook of Agriculture; 1942. p. 1217–25. dium and Haemoproteus mitochondrial DNA amplified from birds. Proc 106. Stabler RM, Kitzmiller NJ, Braun CE. Plasmodium in a Darwin’s tinamou Biol Sci. 2000;276:1583–9. from Colorado. J Parasitol. 1973;59:395. 78. International Commission on Zoological Nomenclature. International 107. Huff CG. Plasmodium hexamerium, n. sp. from the blue-bird, inoculable code of zoological nomenclature. 4th ed. London: The International to canaries. Am J Hyg. 1935;22:274–7. Trust for Zoological Nomenclature; 1999. 108. Novy FG, MacNeal WJ. Trypanosomes and bird malaria. Am Med. 79. Landau I, Chavatte JM, Peters W, Chabaud A. The sub-genera of avian 1904;8:932–4. Plasmodium. Parasite. 2010;17:3–7. 109. Telford SR, Nayar JK, Foster GW, Knight JW. Plasmodium forresteri n. sp., 80. Chavatte JM, Uzbekov R, Paperna I, Richard-Lenoble D, Landau I. from raptors in Florida and Southern Georgia: its distinction from Plas- Ultrastructure of erythrocytic stages of avian Plasmodium spp. of the modium elongatum morphologically within and among host species sub-genus Novyella and its “globule”. Parasite. 2010;17:123–7. and by vector susceptibility. J Parasitol. 1997;83:932–7. 81. Versiani V, Gomes BF. Sobre um novo hematozoário da galinha— 110. Gabaldon A, Ulloa G. A new species of the subgenus Novyella (Hae- “Plasmodium juxtanucleare” n. sp. (Nota prévia). Rev Brasil Biol. mosporina, Plasmodiidae) from Aramides cajanea (Gruiformes, Rallidae). 1941;1:231–3. In: Canning EU, editor. Parasitological topics. A presentation volume 82. Raffaele G II. Plasmodium della civetta (Athene noctua). Riv Malariol. to P.C.C. Garnham, F.R.S. on the occasion of his 80th birthday. Madison: 1931;10:684–8. Society of protozoologists; 1981. p. 100–5. 83. Brumpt E. Paludisme aviaire: Plasmodium gallinaceum n. sp. de la poule 111. Sergent E, Sergent E, Catanei A. Sur un parasite nouveau du paludisme domestique. C R Hebd Séances Acad Sci. 1935;200:783–5. des oiseaux. C R Hebd Séances Acad Sci Paris. 1928;186:809–11. 84. Bano L, Abbasi Z. A new species of avian malaria parasite, Plasmodium 112. Manwell RD. How many species of avian malaria parasites are there? coturnixi, from Coturnix coturnix from Kohat (N.W.F.P., Pakistan). Bull Zool. Am J Trop Med. 1935;15:265–83. 1983;1:17–22. Valkiūnas and Iezhova Malar J (2018) 17:212 Page 24 of 24 113. Manwell RD, Sessler GJ. Plasmodium paranucleophilum n. sp. from a 139. Wolfson F. Plasmodium oti n. sp., a Plasmodium from the eastern screech South American tanager. J Protozool. 1971;18:629–32. owl (Otus asio naevius), infective to canaries. Am J Hyg. 1936;24:94–101. 114. Chagas CRF, Valkiūnas G, Nery CVC, Henrique PC, Gonzalez IHL, Mon- 140. Johnston HT, Cleland JB. Notes on some parasitic Protozoa. Proc Linn teiro EF, et al. Plasmodium (Novyella) nucleophilum from an Egyptian Soc NSW. 1909;34:501–13. Goose in São Paulo Zoo, Brazil: microscopic confirmation and molecular 141. Brumpt E. Paludisme aviaire: Plasmodium paddae n. sp. du calfat (Padda characterization. Int J Parasitol Parasites Wildl. 2013;2:286–91. oryzivora). Utilisation de ce parasite pour les recherches chimiothé- 115. Christensen BM, Barnes HJ, Rowley WA. Vertebrate host specificity and rapiques du paludisme. C R Hebd Séances Acad Sci. 1935;200:967–70. experimental vectors of Plasmodium (Novyella) kempi sp. n. from the 142. Chakravarty M, Kar AB. Studies on Haemosporidia from Indian birds— eastern wild turkey in Iowa. J Wildl Dis. 1983;19:204–13. Series II. Proc Indian Acad Sci. 1945;22:63–9. 116. Carini A. Sur un nouvel hématozoaire du pigeon. C R Hebd Séances 143. Fantham HB, Porter A. On a Plasmodium (Plasmodium relictum var. Mém Soc Biol. 1912;73:396–8. spheniscidae n. var.) observed in four species of penguins. Proc Zool Soc 117. Ilgūnas M, Palinauskas V, Iezhova TA, Valkiūnas G. Molecular and mor- London. 1944;114:279–92. phological characterization of two avian malaria parasites (Haemospor- 144. Mazza S, Fiora A. Proteosoma de mirlo, Planesticus anthracinus (Burm.) y ida: Plasmodiidae), with description of Plasmodium homonucleophilum Leucocytozoon (sic) di Benteveo, Pitangus sulphuratus bolivianus (Latv.) n. sp. Zootaxa. 2013;3666:49. y fueguero Piranga flava ( Viell.) de Tumbaya, Jujuy. 5th Reunion Soc 118. de Jong AC. Plasmodium dissanaikei n. sp. a new avian malaria parasite Argent Patol Reg Norte. 1930;2:993-5. from the rose-ringed parakeet of Ceylon, Psittacula krameri manillensis. 145. Laveran A, Marullaz M. Sur deux hémamibes et un toxoplasme du Ceylon J Med Sci. 1971;20:41–5. Liothrix luteus. Bull Soc Pathol Exot. 1914;7:21–5. 119. Huff CG. Plasmodium elongatum n. sp., an avian malarial organism with 146. Brumpt E. Précis de Parasitologie. Paris: Masson; 1910. an elongate gametocyte. Am J Hyg. 1930;11:385–91. 147. Huang J-C, Huang D-F, Jiang J-B. A new species of the genus Plasmo- 120. Valkiūnas G, Zehtindjiev P, Dimitrov D, Križanauskienė A, Iezhova dium, Plasmodium arachnidi from the domestic pigeon in Guangzhou. TA, Bensch S. Polymerase chain reaction-based identification of Acta Vet Zoot Sinica. 1995;26:352–7. Plasmodium (Huffia) elongatum, with remarks on species identity 148. Huang JC, Huang DF. A new species of the bird malarial parasite—Plas- of haemosporidian lineages deposited in GenBank. Parasitol Res. modium (Novyella) bambusicolai (Sporozoa: Plasmodiidae). Acta Zoot 2008;102:1185–93. Sinica. 1995;20:385–90. 121. Telford SR, Forrester DJ. Plasmodium (Huffia) hermani sp. n. from wild 149. Grés V, Les Landau I, de Lophura Plasmodium. Les Plasmodium de turkeys (Meleagris gallopavo) in Florida. J Protozool. 1975;22:324–8. Lophura (Phasianidae): redescription de P. lophurae Coggeshall, 122. Muniz J, Soares R, Batista S. Sôbre uma espécie de Plasmodium parasita 1938 et description de deux nouvelles espèces. Zoosystema. do Ramphastos toco Müller, 1776. Plasmodium huffi n. sp. Rev Bras 1938;1997(19):545–55. Malariol. 1951;3:339–44. 150. Sarkar AC, Ray HN. A new malarial parasite, Plasmodium (Garnhamella) 123. Wiersch SC, Maier WA, Kampen H. Plasmodium (Haemamoeba) cath- conturnixae n. subgen., n. sp., from black breasted quail, Coturnix emerium gene sequences for phylogenetic analysis of malaria parasites. coromandelica (Aves: Galliformes). In: Progress in protozoology. Abstr. III Parasitol Res. 2005;96:90–4. Inter. Congr. Protozool. Leningrad; 1969. p. 353. 124. Perkins SL, Schall JJ. A molecular phylogeny of malarial parasites recov- 151. Laird M. Avian malaria in the Asian tropical subregion. Singapore: ered from cytochrome b gene sequences. J Parasitol. 2002;88:972–8. Springer; 1998. 125. Omori S, Sato Y, Isobe T, Yukawa M, Murata K. Complete nucleotide 152. Grassi B, Feletti R. Parasites malariques chez les oiseaux. Arch Ital Biol. sequences of the mitochondrial genomes of two avian malaria proto- 1890;13:297–300. zoa, Plasmodium gallinaceum and Plasmodium juxtanucleare. Parasitol 153. Castellani A, Chalmers AJ. Manual of tropical medicine. London: Manual Res. 2007;100:661–4. of tropical Medicine; 1910. 126. Celli A, Sanfelice F. Ueber die Parasiten des rothen Blutkörperchens 154. Stabler RM, Holt PA, Ellison LN. A new malaria from the spruce grouse. J im Menschen und in Thieren. Fortschr Med. 1891;9:499–511, 541–52, Colo-Wyo Acad Sci. 1965;5:49. 581–6. 155. He J-G, Huang J-C. A new species of avian malaria parasite from 127. Gilruth JA, Sweet G, Dodd S. Notes on blood parasites. Proc R Soc Victo- Pycnonotus jocosus (Sporozoa: Plasmodiidae). Acta Zoot Sinica. ria. 1910;23:231–41. 1993;18:129–33. 128. Russell PF. Avian malaria studies, V. Plasmodium capistrani sp. nov., an 156. Oliger IM. New species of parasites of tetraonid birds. Uch zap Chuvash- avian malaria parasite in the Philippines. Philipp J Sci. 1932;48:269–89. skogo gos ped inst. 1956;3:329–35 (in Russian). 129. de Mello IF. Further contribution to the study of blood parasites of 157. Corradetti A, Neri I, Palmieri C, Verolini F, Giuliani V, Scanga M. Note the Indian birds, together with a list of the hemoparasites hitherto su Plasmodium vaughani e su un plasmodio con ciclo schizogonico recorded. J R Asiatic Soc Beng. 1936;2:95–122. endoemoblastico di tipo elongatum rinvenuti in Turdus merula. Paras- 130. de Mello IF. A contribution to the study of the blood parasites of some sitologia. 1961;3:97–100. Indian birds. Proc Indian Acad Sci (Sec. B). 1935;1:349–58. 158. Partsvanidze MI. Plasmodium malariae raupachi from the turkey, new 131. Basu BC. Studies on a malarial infection in a paddy bird. J Malar Inst species. Zakavk vet vestn. 1914;6:86–7 (in Russian). India. 1938;1:273–84. 159. Stabler RM, Datel RJ. First record of malaria in American falcons (Falco 132. Ishiguro H. Plasmodium japonicum, a new species of malaria parasite sparverius). J Colo-Wyo Acad Sci. 1959;4:59. pathogenic for the domestic fowl. Bull Fac Agr Yamaguti Univ. 160. Paperna I, Keong MSC, May CYA. Haematozoan parasites found in birds 1957;8:723–33 (article in Japanese). in peninsular Malaysia, Singapore, Sarawak and Jawa. Raffles Bull Zool. 133. Raffaele G. Considerazioni sulla specificità dei plasmodili. Arch Zool Ital. 2008;56:211–43. 1966;51:273–83. 161. Bray RS. A check-list of the parasitic Protozoa of West Africa with some 134. Labbé A. Recherches zoologiques et biologiques sur les para- notes on their classification. Bull Inst Fr Afr Noir. 1964;26:238–315. sites endoglobulaires du sang des vertébrés. Arch Zool Exp Gen. 162. Yarrington JT, Whitehair CK, Corwin RM. Vitamin E-selenium defi- 1894;2:55–258. ciency and its influence on avian malarial infection in the duck. J Nutr. 135. Raffaele G. Osservazioni sui plasmodidi degli uccelli. Riv Malariol. 1973;103:231–41. 1930;9:209–18. 163. Fantham HB, Porter A. Plasmodium struthionis, sp. n., from Sudanese 136. Corradetti A, Scanga M. Plasmodium (Novyella) vaughani subsp. ostriches and Sarcocystis salvelini, sp. n., from Canadian speckled trout merulae, n. subsp., parassita di Turdus merula, con descrizione del ciclo (Salvelinus fontinalis), together with a record of a Sarcocystis in the eel pre-eritrocitico. Parassitologia. 1972;14:85–93. pout (Zoarces angularis). Proc Zool Soc London. 1943;112:25–30. 137. Das Gupta BM, Siddons LB. On a Plasmodium sp. of the Malay chestnut- 164. Bhaskar Rao TS, Devi A, Bhaskar Rao T. Studies on experimental infec- bellied munia [Munia atricapilla atricapilla ( Vieill)]. Indian Med Gaz. tion of Plasmodium venkataramiahii of the crow Corvus splendens in the 1941;76:148–50. chicks. In: Abstracts. 5th Int Congr Protozool. New York. Abstr. 194; 1977. 138. Laveran A. Sur une Haemamoeba d’une mésange (Parus major). C R Séances Soc Biol Fil. 1902;54:1121–4.
Malaria Journal – Springer Journals
Published: May 29, 2018
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