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Diversity of Haemosporidian Parasites in Mississippi Songbirds
Haley N. Bodden and Diana C. Outlaw

Southeastern Naturalist, Volume 18, Issue 2 (2019): 314–320

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Southeastern Naturalist H.N. Bodden and D.C. Outlaw 2019 Vol. 18, No. 2 314 2019 SOUTHEASTERN NATURALIST 18(2):314–320 Diversity of Haemosporidian Parasites in Mississippi Songbirds Haley N. Bodden1 and Diana C. Outlaw1,* Abstract - Haemosporidian parasites are extremely diverse in birds. The more bird hosts that are tested, the greater the diversity of haemosporidians that is found. Here, we conducted a survey of haemosporidians in the local populations (Oktibbeha County, MS) of common and abundant songbirds. We captured local songbirds with mist nets and collected blood samples from the brachial vein for screening and identification of haemosporidians. Parasite prevalence was 57%, and we detected 3 genera of haemosporidians. We documented 3 Haemoproteus lineages, with 1 novel lineage (3% different than any known lineage); 2 Leucocytozoon lineages were found, neither of which were novel; and 8 Plasmodium lineages were found, one of which was novel (5% different than any known lineage), and 1 of which that may be novel (3% different than any known lineage). We detected Leucocytozoon for the first time in Mississippi songbirds, indicating the importance of surveying for understanding putative haemosporidian range shifts. Introduction Avian malaria has been linked to the huge decline of bird populations in some areas, even leading in some instances to near extinction (Atkinson 2008). Over the evolutionary history of malaria (haemosporidian) parasites, they have spread to over 10,000 avian species, with new haemosporidians being discovered frequently (Daszak et al. 2000, Ricklefs et al. 2014). Climate change has created an opportunity for these parasites to expand to parts of the world they have never been before, primarily due to the distributional changes in the dipertan vectors (see Loiseau et al. 2012, Marzal et al. 2014, Sehgal 2015). Studies have shown how invasive certain species of haemosporidians can be, and how detrimental these parasites are to bird populations (Marzal et al. 2015). The International Union for Conservation of Nature has listed the avian malaria parasite Plasmodium relictum Gilmruth, Sweet, and Dodd in the top 100 worst invasive species in the world. Avian malaria prevalence can vary drastically in different regions (Lauron et al. 2015, Martinsen et al. 2016, Walther et al. 2015), with bird migration increasing the possibility of parasite transmission and potential host switches (Dodge et al. 2013, Ricklefs et al. 2017). More than 325 bird species migrate annually on the Mississippi Flyway and there are numerous permanent residents (Mississippi Chapter, National Audubon Society 2019); thus, the potential for parasite transmission is relevant in this area. We conducted a survey of the haemosporidians in local bird populations over the course of 1 year in order to preliminarily assess the diversity of avian 1Department of Biological Sciences, Mississippi State University, PO Box GY, Mississippi State, MS 39762. *Corresponding author - doutlaw@biology.msstate.edu. Manuscript Editor: Frank Moore Southeastern Naturalist 315 H.N. Bodden and D.C. Outlaw 2019 Vol. 18, No. 2 haemosporidians across multiple, common bird species. We know from previous studies that there is a large number of local birds infected with haemosporidians (Fast et al. 2016, Walstrom and Outlaw 2017). This survey has allowed us to expand our knowledge of infection in a wider range of local passerines. Methods Field work We used mist nets to capture passerines in Starkville, MS, from January 2017 until January 2018. The 3 netting locations are listed in Table 1; we conducted most netting 3–4 d per week, either in the late morning or late afternoon, depending on the availability of student help. We captured 8 passerine species: Cardinalis cardinalis (Northern Cardinal), Baeolophus bicolor (Tufted Titmouse), Carpodacus mexicanus (House Finch), Poecile carolinensis (Carolina Chickadee), Thryothorus ludovicianus (Carolina Wren), Toxostoma rufum (Brown Thrasher), Setophaga petechia (Yellow Warbler), and Troglodytes aedon (House Wren), for a total of 68 birds. We collected blood samples via brachial venipuncture and stored them at -20 °C in RNAlater (Sigma-Aldrich Company, St. Louis, MO). The sample size was limited because ours was a 1-year project involving multiple undergraduate students and funding was limited. DNA extraction and parasite detection We extracted genomic DNA from the blood samples using the DNeasy Blood and Tissue kit (QIAGEN, Inc., Germantown, MD.). We employed the DNeasy protocol “Purification of Total DNA from Animal Blood or Cells” for the nucleated avian blood samples. We detected haemosporidian parasites from the extracted DNA by amplification of the parasite’s mitochondrial cytochrome b (cyt b) gene via the polymerase chain reaction (PCR). We conducted a nested PCR that amplified a specific segment of the parasite’s cyt b gene. The initial PCR used the primers HaemNF1 and HaemNR3 (Hellgren et al. 2004) and amplified Plasmodium, Parahaemoproteus, and Haemoproteus species. We then ran a nested PCR using the Haem PCR as the template. The second set of primers comprised UNIVF and UNIVR1 (Drovetski et al. 2014); these primers amplified the above genera and Leucocytozoon. We performed the initial and nested reactions with the same volume of each reagent the same final volume. The master mix consisted of 2.5 μl Ex Taq Buffer, 2.0 μl dNTP mixture, 0.5 μl BSA, 0.5 μl of each primer, 0.125 μl Ex Taq, 1.0 μl of the DNA template, and 17.875 μl of DI water for each reaction. We used 24 μl of the master mix along with 1μl of the DNA (for the initial PCR) or 1 μl of the initial reaction (for the nested PCR) in each sample. We ran the initial reaction Table 1. Netting locations. Netting location Locality GPS coordinates (°N, °W) South Farm Research Area Starkville, MS 33.420806, 88.783042 Mississippi State University Starkville, MS 33.454135, 88.783455 Critz Street, Starkville MS Starkville, MS 33.471369, 88.811081 Southeastern Naturalist H.N. Bodden and D.C. Outlaw 2019 Vol. 18, No. 2 316 in a thermal cycler under the following conditions: 3 min at 94 °C, 35 cycles of 30 sec at 94 °C, 30 sec at 55 °C, 45 min at 72 °C, 10 min at 72 °C, and a hold at 4 °C. We ran nested reactions in a thermal cycler for 3 min at 94 °C, 41 cycles at 94 °C for 30 sec, 30 sec at 52 °C, 45 sec at 72 °C, 10 min at 72 °C, and held at 4 °C. We visualized the PCRs on a 1% agarose gel via electrophoresis. We used the QIAquick PCR Purification kit (QIAGEN) to clean the positive samples to prepare for sequencing. We sent the cleaned PCR products to the Arizona State University DNA lab (Phoenix, AZ) for sanger sequencing of the forward and reverse strands on an Applied Biosystems 3730 capillary sequencer (ThermoFisher Scientific, Carlsbad, CA). Genus determination Once the sequences were received, we made a consensus sequence by aligning the forward and reverse strands using Sequencher V.5.4.6. We used the NCBI Basic Local Alignment Service Tool (BLAST) to extract either the consensus or the sequence and determine the parasite lineage. Genetic lineage identification We included alignments of this study’s sequences from each genus in a BLAST search of both NCBI and the MalAvi databases (Bensch et al. 2009). We set a 98% sequence similarity as a cut-off for species/lineage identification. We reconstructed phylogenetic trees for each genus for verification of lineage identification (reconstructions not shown) using the cyt b sequences acquired from this study’s sampling as well as those from MalAvi sequences that would provide the most diversity in the phylogeny after an initial inclusion of all known sequences. We aligned all the sequences and checked for gaps or stop codons in Sequencher V.5.4.6. We performed a Bayesian analysis in BEAUTi and BEAST using the following parameters: substitution model = GTR, base frequencies = estimated, clock model = uncorrelated relaxed clock, site heterogeneity model = Gamma + invariant sites, and tree prior-yule process (v.1.8.4; Drummond et al. 2012). Log likelihood stabilization of model parameters was determined in Tracer (v.1.6; Rambaut and Drummond 2007). Results Prevalence We sampled a total of 68 birds from 3 locations in Starkville, MS. We sampled 8 passerine species: Northern Cardinal (45 sampled, 33 infected), Tufted Titmouse (8 sampled, 1 infected), Carolina Chickadee (3 sampled, 2 infected), House Finch (6 sampled, 1 infected), Brown Thrasher (2 sampled, 1 infected), House Wren (1 sampled, 1 infected), Yellow Warbler (2 sampled, 0 infected), Carolina Wren (1 sampled, 0 infected). We determined, via PCR, that the overall parasite prevalence in the samples was 57.4% (Table 2). We detected 3 parasite genera: Plasmodium, Haemoproteus, and Leucocytozoon. Of the 68 total birds sampled, we found that 47.10% of the infections were caused by the genus Plasmodium, 8.80% by Haemoproteus, and 2.9% by Leucocytozoon Southeastern Naturalist 317 H.N. Bodden and D.C. Outlaw 2019 Vol. 18, No. 2 (Table 3). Three samples did not sequence well, and we excluded them from the analysis. All lineages have been deposited in the Dryad Respository under the data package title “Data from: Diversity of haemosporidian parasites in Mississippi songbirds” (DOI:10.5061/dryad.8kd1d3f). Genetic lineage identification We identified 8 lineages for Plasmodium (Table 4), of which 2 were novel. We also detected both Plasmodium circumflexum Kikuth and P. juxtnucleare Versiani and Gomes, as well as what seems to be P. lutzi Lucena. One novel lineage is 95% similar to an Old World lineage found in Europe. We identified 3 lineages for Haemoproteus, (Table 4), and one of these is potentially novel (97% similarity to a known lineage). We identified 2 lineages for Leucocytozoon (Table 4), neither of which is novel, though neither has previously been found in any songbird. Discussion Studies to explore Haemosporidians in Mississippi are recent, but there are several features in common amongst these studies (Fast et al. 2016, Larson et al. 2017, Walstrom and Outlaw 2017). The first is that Mississippi seems to harbor unique parasite lineages that are not found elsewhere. The second is that Mississippi is home to a diverse array of parasites that seemingly come from its prominent position along the Mississippi flyway, one of the largest pathways of birds migrating from north to south, annually. Table 3. Parasite prevalence by haemosporidian genus. Plasmodium Haemoproteus Leucocytozoon Northern Cardinal 28 5 1 Tufted Titmouse 0 0 1 Carolina Chickadee 2 0 0 House Finch 1 0 0 Brown Thrasher 0 1 0 House Wren 1 0 0 Total prevalence % 47.10% 8.80% 2.90% Table 2. Parasite prevalence. Number Number Species collected infected Prevalence Cardinalis cardinalis L. (Northern Cardinal) 45 33 73% Baeolophus bicolor L. (Tufted Titmouse) 8 1 13% Poecile carolinensis (Audubon) (Carolina Chickadee) 3 2 67% Carpodacus mexicanus (Müller) (House Finch) 6 1 17% Toxostoma rufum (L.) (Brown Thrasher) 2 1 50% Troglodytes aedon (Vieillot) (House Wren) 1 1 100% Setophaga petechia (L.) (Yellow Warbler) 2 0 0% Thryothorus ludovicianus (Latham) (Carolina Wren) 1 0 0% Total 68 39 57% Southeastern Naturalist H.N. Bodden and D.C. Outlaw 2019 Vol. 18, No. 2 318 The diversity of Plasmodium parasites we detected is not surprising and is entirely consistent with the previous studies mentioned above, but our detection of 2 Leucocytozoon parasites in songbirds was unexpected. We conducted a survey of waterfowl and detected Leucocytozoon in Aythya affinis (Eyton) (Lesser Scaup) and Phalacrocorax auratus (Lesson), (Double Crested Cormorant) (data not shown). These 2 “infections” may simply be the result of spillover and may not represent viable infections. Our findings do suggest, however, that the vectors of these parasites are taking advantage of multiple bloodmeals, an issue we are currently addressing with research projects involving Simuliidae (blackflies) and Culicinae and Anophelinae (mosquitoes). Surveys of local haemosporidians are crucial in monitoring efforts. Haemosporidians are expanding their geographic range and acquiring new hosts in new areas, particularly along migratory pathways (Ricklefs et al. 2017). This information reinforces that our knowledge of the diversity in haemosporidians remains extremely limited (see also Valikunas 2005). Future directions One result from comparing 2 previous projects on Northern Cardinals and Tufted Titmice (Fast et al. 2016, Walstrom and Outlaw 2017), was the difference between which parasites infected each host species. Plasmodium seemed to prefer Tufted Titmice to Northern Cardinals and Parahaemoproteus seemed to prefer Northern Cardinals. The samples from this study, combined with our previously collected samples, will allow us to conduct natural differential gene expression experiments between host species to compare the immune responses to different hosts to both the same and different parasites. Table 4. Parasite lineage identification. Putative novel lineages are noted with an asterisk ( *). Number of Species (if known) and/or Lineage times found % similarity to MalAvi Lineage (name) Haemoproteus H1 4 98–100 (SIAMEX01) H2 1 99 H3* 1 97 Leucocytozoon L1 1 100 (AIXGAL01) L2 1 98 Plasmodium P1 2 Plasmodium circumflexum/98–99 P2 4 100 (FIPAR01) P3 7 98–100 (TRICRI01) P4 7 P. juxtanucleare/99–100 P5 3 P. lutzi P6 1 98 P7* 1 95 P8* 1 97 Southeastern Naturalist 319 H.N. Bodden and D.C. Outlaw 2019 Vol. 18, No. 2 Acknowledgments Funding for the project was provided by the National Institutes of Health R03AI117223- 01A1 to D.C. Outlaw. 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