Parasitaemia data and molecular characterization of Haemoproteus catharti from New World vultures (Cathartidae) reveals a novel clade of Haemosporida
- Michael J. Yabsley1, 2Email authorView ORCID ID profile,
- Ralph E. T. Vanstreels3, 4,
- Ellen S. Martinsen5, 6,
- Alexandra G. Wickson1,
- Amanda E. Holland1, 7,
- Sonia M. Hernandez1, 2,
- Alec T. Thompson1,
- Susan L. Perkins8,
- Christopher J. West9,
- A. Lawrence Bryan7,
- Christopher A. Cleveland1, 2,
- Emily Jolly1,
- Justin D. Brown10,
- Dave McRuer11,
- Shannon Behmke12 and
- James C. Beasley1, 7
© The Author(s) 2018
Received: 31 October 2017
Accepted: 25 December 2017
Published: 8 January 2018
New World vultures (Cathartiformes: Cathartidae) are obligate scavengers comprised of seven species in five genera throughout the Americas. Of these, turkey vultures (Cathartes aura) and black vultures (Coragyps atratus) are the most widespread and, although ecologically similar, have evolved differences in morphology, physiology, and behaviour. Three species of haemosporidians have been reported in New World vultures to date: Haemoproteus catharti, Leucocytozoon toddi and Plasmodium elongatum, although few studies have investigated haemosporidian parasites in this important group of species. In this study, morphological and molecular methods were used to investigate the epidemiology and molecular biology of haemosporidian parasites of New World vultures in North America.
Blood and/or tissue samples were obtained from 162 turkey vultures and 95 black vultures in six states of the USA. Parasites were identified based on their morphology in blood smears, and sequences of the mitochondrial cytochrome b and nuclear adenylosuccinate lyase genes were obtained for molecular characterization.
No parasites were detected in black vultures, whereas 24% of turkey vultures across all sampling locations were positive for H. catharti by blood smear analysis and/or PCR testing. The phylogenetic analysis of cytochrome b gene sequences revealed that H. catharti is closely related to MYCAMH1, a yet unidentified haemosporidian from wood storks (Mycteria americana) in southeastern USA and northern Brazil. Haemoproteus catharti and MYCAMH1 represent a clade that is unmistakably separate from all other Haemoproteus spp., being most closely related to Haemocystidium spp. from reptiles and to Plasmodium spp. from birds and reptiles.
Haemoproteus catharti is a widely-distributed parasite of turkey vultures in North America that is evolutionarily distinct from other haemosporidian parasites. These results reveal that the genetic diversity and evolutionary relationships of avian haemosporidians are still being uncovered, and future studies combining a comprehensive evaluation of morphological and life cycle characteristics with the analysis of multiple nuclear and mitochondrial genes will be useful to redefine the genus boundaries of these parasites and to re-evaluate the relationships amongst haemosporidians of birds, reptiles and mammals.
The Order Haemosporida contains numerous vector-borne protozoan blood parasites of reptilian, avian and mammalian hosts [1, 2]. These parasites utilize a wide range of vectors and are found on all continents except Antarctica. There are numerous genera of haemosporidian parasites, four of which have species recorded in avian hosts: Plasmodium (described in 1885), Haemoproteus (1890), Leucocytozoon (1904) and Fallisia (1974) . Considering genetic evidence and differences in natural history, however, many researchers have suggested that the two subgenera of Haemoproteus (Haemoproteus and Parahaemoproteus) and the two subgenera of Leucocytozoon (Akiba and Leucocytozoon) should be elevated to genera [3, 4]. A recent study has also uncovered genetic evidence that indicates Haemoproteus antigonis, a parasite from cranes (Gruidae), represents a novel clade that is paraphyletic with other known Haemosporida, potentially meriting a separate genus .
Despite the vast amount of information on the haemosporidians of birds, knowledge about the species that infect vultures is still very limited. Vultures are large, obligate scavenging birds that are divided into two families, the Old World vultures (Accipitridae: Aegypiinae, Gypaetinae) and the New World vultures (Cathartidae). The morphological and physiological similarities between these groups are a remarkable example of convergent evolution [6, 7]. There are currently seven species of New World vultures in five genera, four of which are monotypic . Of these, turkey vultures (Cathartes aura, TUVU) and black vultures (Coragyps atratus, BLVU) are the most widespread, ranging from southern South America into the United States, and even Canada in the case of turkey vultures. Despite similarities in appearance and functional role in ecosystems, black and turkey vultures have evolved unique morphological, physiological, and behavioural differences that result in interspecific differences in foraging behaviour [9, 10]. With the exception of the California condor (Gymnogyps californianus), which has a restricted distribution in the western United States, the remaining New World vulture species reside in Central and South America .
Published records of hemosporidian parasites in New World vultures
Turkey vulture (TUVU)
USA (Washington DC)
USA (Washington DC)
USA (Maryland and New Jersey)
Haemoproteus catharti and Plasmodium sp.c
USA (South Carolina)
Black vulture (BLVU)
Plasmodium elongatum and Leucocytozoon toddi
Brazil (São Paulo)
In this study, morphological and molecular methods were used to investigate the epidemiology and evolution of haemosporidian parasites of New World vultures sampled at six states of the USA. These data challenge the placement of H. catharti in the genus Haemoproteus, and instead suggest that these parasites represent a novel evolutionary lineage of haemosporidians, possibly meriting a separate genus.
Results of haemosporidian detection on New World vultures from the United States
Turkey vulture (TUVU)
Blood, muscle, and spleen
Blood, muscle and spleen
Black vulture (BLVU)
Blood, muscle, spleen
In Georgia, additional four TUVU and three BLVU samples were obtained from nuisance removals in Burke County (33°04′59″ N 82°01′19″ W). In Virginia, samples were obtained from two injured TUVU admitted for rehabilitation (Albemarle and Franklin counties) (38°01′43″ N 78°29′06″ W and 37°00′02″ N 79°53′05″ W) and from 10 TUVU sampled during a study on lead exposure in Chesterfield county (37°22′35″ N 77°30′23″ W). Additionally, two BLVU from Bedford and Louisa counties admitted for rehabilitation were sampled in Virginia (37°19′53″ N 79°31′23″ W and 38°01′31″ N 78°00′11″ W). In Florida, samples from two TUVU were obtained from birds from Lee County admitted for rehabilitation (26°38′35″ N 81°52′07″ W and 26°33′52″ N 81°57′53″ W). In Pennsylvania, samples were collected from three TUVU found dead in Franklin, Centre and Huntingdon counties (39°55′58″ N 77°39′05″ W, 40°46′30″ N 77°52′51″ W, and 40°28′39″ N 78°01′05″ W, respectively) that were submitted for diagnostic evaluation. Collection of the remainder of these samples were reviewed and approved by UGA’s IACUC protocol A2014 10-018.
When possible, blood samples were collected from the metatarsal vein. Two thin blood smears were immediately prepared in the field, air dried, fixed in 100% methanol, and later taken back to the laboratory to be stained with a modified Giemsa (DipQuick, Jorgensen Laboratories, Inc., Loveland, CO, USA). Remaining blood from most sites was preserved in heparin and frozen until PCR analysis. Blood samples from TUVU from California were placed on filter paper, dried and stored in a desiccant until testing. For deceased vultures, clotted blood, muscle, and spleen samples were obtained and frozen for PCR analysis. See Table 2 for which samples were collected from the different groups of birds.
Blood smears were examined at 1000× under oil immersion to determine infection status for blood parasites, with a minimum of 20,000 examined erythrocytes. Parasites were morphologically identified based on descriptions in the literature [2, 11, 16, 20]. Parasitaemia (no. parasites/erythrocytes examined) and the following morphometric parameters were obtained for a subset of mature gametocytes : length and width of the parasite, length and width of infected and uninfected erythrocytes, number of pigment granules in the parasite, position of the parasite within the erythrocyte, and the ratio of nuclear displacement.
All tissue and/or blood samples were tested for Haemoproteus and Plasmodium using nested polymerase chain reaction (PCR). DNA was extracted from 10 µL of blood or ~ 10 mg of tissue using a commercial kit per the manufacturer’s instructions (Qiagen Dneasy Blood & Tissue Kit, Germantown, MD, USA). Nested PCR targeting the mitochondrial cytochrome b (cytb) gene was conducted as described using primary primers HaemNFI and HaemNR3 and nested primers HaemF and HaemR2 . A subset of TUVU samples from Virginia (n = 12), Georgia (n = 3), Pennsylvania (n = 3) and Florida (n = 2) were tested for Leucocytozoon sp. using nested PCR using primary primers HaemNFI and HaemNR3 and nested primers HaemFL and HaemR2L . Amplification products were visualized in 2% agarose gels stained with GelRed (Biotium, Hayward, CA, USA).
Amplification products of 18 TUVU samples (South Carolina = 11, Virginia = 3, Florida = 2, Pennsylvania = 1, California = 1) were extracted from the gel, purified using the QIAquick gel extraction kit (Qiagen), and submitted for bi-directional sequencing (using the HaemF and HaemR2 primers; 479 bp fragment) at the Georgia Genomics Facility (Athens, GA, USA). For one TUVU sample from South Carolina, a longer region of the cytb gene (725 bp) was sequenced using primers DW2 and DW4 . For two TUVU samples from South Carolina, the nuclear adenylosuccinate lyase (asl) gene was sequenced using the primers described by Martinsen et al. . Sequences obtained in this study were deposited in GenBank (accession numbers MF953291–MF953293).
Phylogenetic analysis of the cytb gene sequences was conducted to compare sequences obtained in this study to those of avian haemosporidians from the MalAvi database , as well as publicly-available sequences of reptilian and mammalian Haemosporida from GenBank (Additional file 1). Sequences were aligned using ClustalW  as implemented in MEGA 6.06 . Bayesian phylogenetic trees were produced using MrBayes 3.2.6 ; the GTR + I + Γ model of nucleotide evolution was used as recommended by jModelTest 2.1.10 . Two Markov chains were run simultaneously for 10 million generations with sampling every 1000 generations; the first 2500 trees (25%) were discarded as a burn-in step and the remaining trees were used to calculate the posterior probabilities.
Comparison of prevalence based on PCR testing between vulture species, sampling sites, sampling method (live capture and nuisance removals vs. those that were in rehabilitation or were found dead), months and years was conducted using Fisher’s exact tests to account for low samples sizes for some groups. Variation of parasitaemia was compared in relation to age group (as determined in ), year and month of sampling, and sampling method was tested using Student’s T test or analysis of variance (ANOVA). Significance level was 0.05.
No parasites were detected by blood smears or PCR testing in BLVU samples. A total of 39/161 (24%) TUVU from six states were positive for haemosporidians by blood smear analysis and/or PCR testing (Table 2). All TUVU tested for Leucocytozoon spp. were PCR negative. The only other blood parasite observed in blood smears were microfilariae detected in two TUVU from South Carolina.
Morphometrics of Haemoproteus catharti as described by Greiner et al.  and in this study
Greiner et al. 
Greiner et al. 
(n = 35)
(n = 20)
(n = 43)
(n = 3)
Gametocyte length (μm)
14.88 ± 1.60
15.4 ± 1.1
13.61 ± 1.30
14.5 ± 0.6
Gametocyte width (μm)
3.6 ± 0.5
3.4 ± 0.5
3.77 ± 0.70
3.7 ± 0.6
Nuclear displacement ratio (NDR)
0.57 ± 0.20
0.59 ± 0.2
0.46 ± 0.10
0.5 ± 0.2
Infected erythrocyte length (μm)
14.6 ± 0.7
15.4 ± 1.0
14.3 ± 0.9
15.2 ± 1.3
Infected erythrocyte width (μm)
8.1 ± 0.8
7.7 ± 0.7
8.2 ± 1.0
7.8 ± 0.8
Pigment granule number
25.6 ± 6.7
24.4 ± 7.0
22 ± 5.4
18.0 ± 7.0
Parasite in contact with host cell nucleus?
55% yes, 45% no
49% yes, 51% no
The apparent prevalence of H. catharti in TUVU was significantly higher in the five eastern states (38/116, 33%) compared to the western state of California (1/45, 2%) (p < 0.001). At the South Carolina study site, the apparent prevalence of H. catharti was higher in 2013 (16/38, 42%) compared to 2014 (12/58, 21%) (p = 0.038) but no difference was noted between month of sampling (April 0/1 (0%) positive, May 10/27 (37%), June 11/42 (21%), and July 7/15 (47%); p > 0.05). For the TUVU sampled in the eastern United States, there was no difference in prevalence by sampling method (live capture/nuisance 35/109, 32% vs. sick or dead birds 4/7, 57%). In blood smear-positive birds (n = 28), parasitaemia was generally low (0.0302 ± 0.0326 parasites/erythrocyte, range 0.0049–0.1094) and was not significantly different in relation to age group (3 juveniles, 25 adults; p = 0.775), year (16 birds sampled in 2013, 12 in 2014; p = 0.369) or month of sampling (10 birds positive in May, 11 in June, 7 in July; p = 0.358).
Partial asl sequences (246 bp) from two TUVU from the South Carolina site were identical. These sequences were nearly equally similar to several avian, mammalian, and reptilian representatives in genera such as Polychromophilus (80–83%), Nycteria (79–84%), and Plasmodium (79–83%). However, the asl gene sequences had much lower identity to the other two avian malaria genera Haemoproteus (76–80%) and Leucocytozoon (71–79%).
Haemoproteus catharti was detected in TUVU at all studied US states, but no evidence of infection was detected in BLVU by either PCR assay or blood smear analysis. This difference is surprising considering that these species have a shared evolutionary history  and have extensive similarities in the functional ecology, as well as in morphological, physiological, and behavioural attributes [9, 10]. The closest relatives of TUVU are endemic to South America, the lesser yellow-headed vulture (Cathartes burrovianus) and the greater yellow-headed vulture (Cathartes melambrotus) . Neither of these species have been examined for blood parasites, thus future studies would be valuable to determine if these species have parasites related to H. catharti. The only haemoparasites detected in the current study were H. catharti and unidentified microfilaria, but paired blood smears and blood for PCR testing were only available from one site so it is possible that other parasites were not detected as the PCR assay used will amplify both Plasmodium and Haemoproteus and lack of amplification of some parasite species with commonly used PCR assays has been reported .
A single nucleotide difference in the cytb gene region was detected in the sequences of H. catharti detected in the two subspecies of TUVU sampled, Cathartes aura septentrionalis from the eastern United States and Cathartes aura meridionalis in California. These two TUVU subspecies do not overlap in residential range and have distinct migration patterns with C. a. septentrionalis being much less migratory than C. a. meridionalis [30, 31]. It is unknown if this genetic difference is due to TUVU subspecies isolation or general geographic variation. In the 1960s, Galindo and Sousa  reported Haemoproteus in TUVU from Panama, which could possibly be in Cathartes aura ruficolis. However, the authors noted that the birds sampled were migrants flying through so it is unknown if infections were actually in migrating C. a. meridionalis from the western United States. Future molecular characterization studies of additional H. catharti samples from the different TUVU subspecies are needed to determine if the parasites vary by subspecies or geographic location.
The phylogenetic analysis of cytb gene sequences revealed that H. catharti is closely related to Haemosporida sp. MYCAMH1, a yet unidentified parasite of wood storks (Mycteria americana) in southeastern USA (Georgia state) and northern Brazil (Amapá state) . Initially MYCAMH1 had been classified as a Haemoproteus sp. and although the morphology of this lineage has not been described, Villar et al.  provided a photomicrograph of a gametocyte that shows the parasite contains pigmented granules and thus generally conforms to Haemoproteus. However, H. catharti and MYCAMH1 constitute a clade that is unmistakably separate from all other Haemoproteus spp., being most closely related to Haemocystidium spp. from reptiles and to Plasmodium spp. from birds and reptiles. The partial asl gene sequence obtained in the current study also suggests that H. catharti is clearly distinct from all other Haemoproteus spp., being instead most similar to other haemosporidian genera, such as Polychromophilus, Nycteria and Plasmodium.
Based on their natural history and morphological characteristics, pigmented haemosporidian that infect avian erythrocytes without forming erythrocytic meronts would traditionally be placed in the genus Haemoproteus . However, the genetic evidence produced in this study suggests this parasite (along with MYCAMH1) might represent a novel genus. This would place H. catharti in an analogous condition to Haemoproteus antigonis, which was recently discovered to represent a separate clade from the remainder Haemoproteus spp. . Thus, it is clear that a taxonomic revision of avian haemosporidians is warranted, possibly with the designation of novel, separate, genera for H. antigonis and H. catharti. It is worth noting that, based on the presence of pigment in gametocytes and the absence of merogony in blood cells, H. antigonis and H. catharti would still be classified in the family Haemoproteidae alongside with other genera such as Haemocystidium, Haemoproteus, Hepatocystis, and Polychromophilus .
There are a number of records of Haemoproteus sp. in BLVU and TUVU for which the parasites were not morphologically or genetically characterized (Table 1); it is reasonable to suspect that some of these records—or perhaps all of them—correspond to H. catharti. Neither of the two species of Haemoproteus recorded in Old World vultures, H. elani and H. janovyi, have been molecularly characterized; therefore, it is not possible at present to evaluate their phylogenetic relationship to H. catharti. Of these, H. elani bears remarkable morphological similarities to H. catharti, as in both species: (a) fully grown gametocytes are halteridial and do not completely encircle the infected erythrocyte nucleus (Fig. 2E–L), (b) fully grown gametocytes have variable contact with the infected erythrocyte nucleus (Fig. 2E, I–K), and (c) fully grown gametocytes fill the infected erythrocytes up to their poles (Fig. 2G) [2, 16]. It is worth noting that H. elani is traditionally considered a parasite of hawks and eagles (Accipitriformes)  and there are only two records of this parasite in an Old World vultures: a lappet-faced vulture (Torgos tracheliotos) captive at the Oklahoma Zoo, USA  and a white-backed vulture (Gyps africanus) sampled at Nossob Camp, Cape of Good Hope, Western Cape, South Africa (IRCAH accession number 103937—M. Bryant, pers. comm.) . Considering the morphological similarities between H. elani and H. catharti, future studies would benefit from evaluating the gene sequences of these parasites to appraise their evolutionary relationship and their relation to H. elani strains from hawks and eagles.
It is interesting to note that the phylogenetic proximity between H. catharti and MYCAMH1 seems to parallel early, and apparently incorrect, suggestions of a close relationship between New World vultures (Cathartiidae) and storks (Ciconiidae) [36, 37]. In recent years, however, it has been proposed that New World vultures and storks have a paraphyletic origin and Cathartiidae should be placed in its own order, Cathartiformes , being most closely related to hawks and eagles . Future studies on the molecular biology of haemosporidian parasites from Accipitridae, Cathartiidae and Ciconiidae would thus be valuable in clarifying the host specificity of these organisms and their transmission and evolution across host taxonomic boundaries.
Lastly, if H. catharti and MYCAMH1 are not closely related to other Haemoproteus spp., it is possible that different vectors are involved in their transmission. Haemoproteus (Parahaemoproteus) spp. and Leucocytozoon caulleryi are transmitted by Ceratopogonidae (biting midges), Haemoproteus (Haemoproteus) spp. are transmitted by Hippoboscidae (louse flies), Plasmodium spp. are transmitted by Culicidae (mosquitoes), and Leucocytozoon (Leucocytozoon) spp. are transmitted by Simuliidae (black flies) . The vectors of the reptile-infecting Haemocystidium spp. (formerly classified as Haemoproteus) are poorly understood but one Haemocystidium species, H. metchnikovi, has been successfully transmitted to painted turtles (Chrysemys picta) by Tabanidae (horse flies) . The vector(s) of Fallisia neotropicalis are unknown but there are some experimental data that suggest mosquitoes may be vectors . It is, therefore, reasonable to consider these families of dipteran insects as potential candidates to be the vectors of H. catharti (and likely MYCAMH1).
Haemoproteus catharti is a widely distributed parasite of TUVU in North America that is evolutionarily distinct from other haemosporidian parasites. These data, alongside those of a recent study on the haemosporidian parasites of North American cranes , reveal that the genetic diversity and evolutionary relationships of avian haemosporidians are still being uncovered. Future studies combining a comprehensive evaluation of morphological and life cycle characteristics with the analysis of multiple nuclear and mitochondrial genes are needed to redefine the genus boundaries of these parasites and to re-evaluate the relationships amongst haemosporidians of birds, reptiles and mammals.
MJY, JCB, SMH designed the study, conducted field work and collected the samples. ESM, AEH, CJW, ALB, JDB, DM, SB, JCB also collected samples. MJY, ESM, SLP, ATT, AGW, EJ, CAC performed the laboratory work. MJY, RETV, ESM, AGW, JCB analyzed the data. RETV, MJY, JCB and SMH wrote the paper. All authors read and approved the final manuscript.
We thank personnel at the Savannah River Site, the Southeastern Cooperative Wildlife Disease Study, and the United States Department of Agriculture for field and lab assistance. We are grateful to Heather Barron, Todd Katzner and Mal Bryant for their valuable contributions of samples.
The authors declare that they have no competing interests.
Availability of data and materials
Gene sequences produced in this study are publicly-available through GenBank (https://0-www-ncbi-nlm-nih-gov.brum.beds.ac.uk/genbank/) under accession numbers MF953291–MF953293. Accession numbers of gene sequences in the phylogenetic analyses are provided in the Additional files.
Ethics approval and consent to participate
Research procedures were reviewed and approved by the Institutional Animal Care and Use Committee of the University of Georgia (A2014 11-010, A2013 01-005) and by the Humboldt State University Institutional Animal Care and Use Committee (08/09.W.89.A).
We acknowledge the National Science Foundation-funded Population Biology of Infectious Diseases program at the University of Georgia for support of AGW and ATT and the National Institute of Health-funded UGA PREP program for partial support of ATT. Additional funding was provided by USDA-APHIS-WS/NWRC, USDOT/FAA, U.S. Fish and Wildlife Service Tribal Wildlife Grants Program and Bureau of Indian Affairs Endangered Species Program, and the U.S. Department of Energy (Award Number DE-FC09-07SR22506) and the sponsorship of the Southeastern Cooperative Wildlife Disease Study by the fish and wildlife agencies of Alabama, Arkansas, Florida, Georgia, Kentucky, Kansas, Louisiana, Maryland, Mississippi, Missouri, Nebraska, North Carolina, Ohio, Oklahoma, Pennsylvania, South Carolina, Tennessee, Virginia, and West Virginia, USA. Support from the states to SCWDS was provided in part by the Federal Aid to Wildlife Restoration Act (50 Stat. 917).
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
- Davies A, Johnston M. The biology of some intraerythrocytic parasites of fishes, amphibia and reptiles. Adv Parasitol. 2000;45:1–107.View ArticlePubMedGoogle Scholar
- Valkiūnas G. Avian malaria parasites and other haemosporidia. Boca Raton: CRC Press; 2005.Google Scholar
- Bennett G, Peirce M. Morphological form in the avian Haemoproteidae and an annotated checklist of the genus Haemoproteus Kruse, 1890. J Nat Hist. 1988;22:1683–96.View ArticleGoogle Scholar
- Bennett G, Peirce M, Ashford R. Avian haematozoa: mortality and pathogenicity. J Nat Hist. 1993;27:993–1001.View ArticleGoogle Scholar
- Bertram MR, Hamer SA, Hartup BK, Snowden KF, Medeiros MC, Outlaw DC, et al. A novel Haemosporida clade at the rank of genus in North American cranes (Aves: Gruiformes). Mol Phylogenet Evol. 2017;109:73–9.View ArticlePubMedGoogle Scholar
- Campbell MO. A fascinating example for convergent evolution: endangered vultures. J Biodivers Endanger Species. 2014;2:132.Google Scholar
- Chung O, Jin S, Cho YS, Lim J, Kim H, Jho S, et al. The first whole genome and transcriptome of the cinereous vulture reveals adaptation in the gastric and immune defense systems and possible convergent evolution between the Old and New World vultures. Genome Biol. 2015;16:215.View ArticlePubMedPubMed CentralGoogle Scholar
- Houston D. New World Vultures (Cathartidae). In: del Hoyo J, Elliott A, Sargatal J, Christie D, de Juana E (editors). Handb. Birds World Alive. Barcelona: Lynx Edicions; 2017; http://www.hbw.com/node/52211 Accessed 2017 Aug 30.
- Wenzel BM, Sieck MH. Olfactory perception and bulbar electrical activity in several avian species. Physiol Behav. 1972;9:287–93.View ArticlePubMedGoogle Scholar
- Turner KL, Abernethy EF, Mike Conner L, Rhodes OE, Beasley JC. Abiotic and biotic factors modulate carrion fate and vertebrate scavenging communities. Ecology. 2017;98:2413–24.View ArticlePubMedGoogle Scholar
- Greiner EC, Mundy PJ. Hematozoa from southern African vultures, with a description of Haemoproteus janovyi sp. n. J Parasitol. 1979;65:147–53.View ArticlePubMedGoogle Scholar
- Halpern N, Bennett GF. Haemoproteus and Leucocytozoon infections in birds of the Oklahoma City Zoo. J Wildl Dis. 1983;19:330–2.View ArticlePubMedGoogle Scholar
- Earlé R, Bennett G, Du Toit H, De Swardt D, Herholdt J. Regional and seasonal distribution of avian blood parasites from northern South Africa. South Afr J Wildl Res. 1991;21:47–53.Google Scholar
- Peirce M, Anderson M. Annual prevalence of haematozoa in White-backed Vulture Gyps africanus nestlings at Kimberley. S Afr Ostrich. 2010;81:269–70.View ArticleGoogle Scholar
- Forrester DJ, Spalding MG. Parasites and diseases of wild birds in Florida. Gainesville: University Press of Florida; 2003.Google Scholar
- Greiner EC, Fedynich AM, Webb SL, DeVault TL, Rhodes OE Jr. Hematozoa and a new haemoproteid species from Cathartidae (New World Vulture) in South Carolina. J Parasitol. 2011;97:1137–9.View ArticlePubMedGoogle Scholar
- Chagas CRF, Valkiūnas G, Guimarães LO, Monteiro EF, Guida FJV, Simões RF, et al. Diversity and distribution of avian malaria and related haemosporidian parasites in captive birds from a Brazilian megalopolis. Malar J. 2017;16:83.View ArticlePubMedPubMed CentralGoogle Scholar
- Holland AE, Byrne ME, Bryan AL, DeVault TL, Rhodes OE, Beasley JC. Fine-scale assessment of home ranges and activity patterns for resident black vultures (Coragyps atratus) and turkey vultures (Cathartes aura). PLoS ONE. 2017;12:e0179819.View ArticlePubMedPubMed CentralGoogle Scholar
- West C, Wolfe J, Wiegardt A, Williams-Claussen T. Feasibility of California Condor recovery in northern California, USA: contaminants in surrogate Turkey vultures and Common Ravens. Condor. 2017;119:720–31.View ArticleGoogle Scholar
- Peirce M, Bennett G, Bishop M. The haemoproteids of the avian order Falconiformes. J Nat Hist. 1990;24:1091–100.View ArticleGoogle Scholar
- Hellgren O, Waldenström J, Bensch S. A new PCR assay for simultaneous studies of Leucocytozoon, Plasmodium, and Haemoproteus from avian blood. J Parasitol. 2004;90:797–802.View ArticlePubMedGoogle Scholar
- Perkins SL, Schall J. A molecular phylogeny of malarial parasites recovered from cytochrome b gene sequences. J Parasitol. 2002;88:972–8.View ArticlePubMedGoogle Scholar
- Martinsen ES, Perkins SL, Schall JJ. A three-genome phylogeny of malaria parasites (Plasmodium and closely related genera): evolution of life-history traits and host switches. Mol Phylogenet Evol. 2008;47:261–73.View ArticlePubMedGoogle Scholar
- Bensch S, Hellgren O, Pérez-Tris J. MalAvi: a public database of malaria parasites and related haemosporidians in avian hosts based on mitochondrial cytochrome b lineages. Mol Ecol Resour. 2009;9:1353–8.View ArticlePubMedGoogle Scholar
- Thompson JD, Gibson T, Higgins DG. Multiple sequence alignment using ClustalW and ClustalX. Curr Protoc Bioinform. 2002, https://doi.org/10.1002/0471250953.bi0203s00.
- Tamura K, Stecher G, Peterson D, Filipski A, Kumar S. MEGA6: molecular evolutionary genetics analysis version 6.0. Mol Biol Evol. 2013;30:2725–9.View ArticlePubMedPubMed CentralGoogle Scholar
- Ronquist F, Huelsenbeck JP. MrBayes 3: bayesian phylogenetic inference under mixed models. Bioinformatics. 2003;19:1572–4.View ArticlePubMedGoogle Scholar
- Darriba D, Taboada GL, Doallo R, Posada D. jModelTest 2: more models, new heuristics and parallel computing. Nat Methods. 2012;9:772.View ArticlePubMedPubMed CentralGoogle Scholar
- Johnson JA, Brown JW, Fuchs J, Mindell DP. Multi-locus phylogenetic inference among New World vultures (Aves: Cathartidae). Mol Phylogenet Evol. 2016;105:193–9.View ArticlePubMedGoogle Scholar
- Amadon D. Notes on the taxonomy of vultures. Condor. 1977;79:413–6.View ArticleGoogle Scholar
- Dodge S, Bohrer G, Bildstein K, Davidson SC, Weinzierl R, Bechard MJ, et al. Environmental drivers of variability in the movement ecology of turkey vultures (Cathartes aura) in North and South America. Phil Trans R Soc B Biol Sci. 2014;369:20130195.View ArticleGoogle Scholar
- Galindo P, Sousa O. Blood parasites of birds from Almirante, Panama with ecological notes on the hosts. Rev Biol Trop. 1966;14:27–46.Google Scholar
- Villar CM, Bryan AL Jr, Lance SL, Braga EM, Congrains C, Del Lama SN. Blood parasites in nestlings of wood stork populations from three regions of the American continent. J Parasitol. 2013;99:522–7.View ArticlePubMedGoogle Scholar
- Martinsen ES, Perkins SL. The diversity of Plasmodium and other Haemosporidians: The interesection of taxonomy, phylogenetics and genomics. In: Carlton JM, Perkins SL, Deitsch KW, editors. Malaria Parasites: Comparative Genomics, Evolution and Molecular Biology. Norfolk: Caister Academic Press; 2013. p. 1–15.Google Scholar
- Bennett G, Earlé R, Du Toit H. A host-parasite catalogue of the haematozoa of the sub-Saharan birds. Onderstepoort J Vet Res. 1992;59:1–73.PubMedGoogle Scholar
- Ligon JD. Relationships of the cathartid vultures. Occas Pap Mus Zool. 1967;651:1–28.Google Scholar
- Avise JC, Nelson WS, Sibley CG. DNA sequence support for a close phylogenetic relationship between some storks and New World vultures. Proc Natl Acad Sci USA. 1994;91:5173–7.View ArticlePubMedPubMed CentralGoogle Scholar
- Chesser RT, Burns KJ, Cicero C, Dunn JL, Kratter AW, Lovette IJ, et al. Fifty-seventh supplement to the American ornithologists’ Union check-list of North American Birds. Auk. 2016;133:544–60.View ArticleGoogle Scholar
- Prum RO, Berv JS, Dornburg A, Field DJ, Townsend JP, Lemmon EM, et al. A comprehensive phylogeny of birds (Aves) using targeted next-generation DNA sequencing. Nature. 2015;526:569.View ArticlePubMedGoogle Scholar
- DeGiusti D, Sterling C, Dobrzechowski D. Transmission of the chelonian haemoproteid Haemoproteus metchnikovi by a tabanid fly Chrysops callidus. Nature. 1973;242:50–1.View ArticlePubMedGoogle Scholar
- Darling S. Some blood parasites (Haemoproteus and Haemogregarina). Bull Soc Path Exot. 1912;5:71–3.Google Scholar
- Wetmore PW. Blood parasites of birds of the District of Columbia and Patuxent Research Refuge vicinity. J Parasitol. 1941;27:379–93.View ArticleGoogle Scholar
- Love GJ, Wilkin SA, Goodwin MH. Incidence of blood parasites in birds collected in southwestern Georgia. J Parasitol. 1953;39:52–7.View ArticlePubMedGoogle Scholar
- Williams NA, Bennett GF. Hematozoa of some birds of New Jersey and Maryland. Can J Zool. 1978;56:596–603.View ArticlePubMedGoogle Scholar
- Webb SL, Fedynich AM, Yeltatzie SK, De Vault TL, Rhodes OE Jr. Survey of blood parasites in black vultures and turkey vultures from South Carolina. Southeast Nat. 2005;4:355–60.View ArticleGoogle Scholar
- Wahl M. Blood-borne parasites in the Black Vulture Coragyps atratus in northwestern Costa Rica. Vulture News. 2013;64:21–30.Google Scholar