Haemocystidium Spp., a Species Complex Infecting Ancient Aquatic

Home , Aconoidasida, Adeleorina, Agamococcidiorida, Arrau turtle, Cnemidophorus gramivagus, Colpodellida

IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Leydy Paola González Camacho

Universidad Nacional de Colombia Facultad de ciencias, Instituto de Biotecnología IBUN Bogotá, Colombia 2019

IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Leydy Paola González Camacho

Tesis o trabajo de investigación presentada(o) como requisito parcial para optar al título de: Magister en Microbiología.

Director (a): Ph.D MSc Nubia Estela Matta Camacho Codirector (a): Ph.D MSc Mario Vargas-Ramírez

Línea de Investigación: Biología molecular de agentes infecciosos Grupo de Investigación: Caracterización inmunológica y genética

Universidad Nacional de Colombia Facultad de ciencias, Instituto de biotecnología (IBUN) Bogotá, Colombia 2019

IV IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

A mis padres, A mi familia, A mi hijo, inspiración en mi vida

Agradecimientos

Quiero agradecer especialmente a mis padres por su contribución en tiempo y recursos, así como su apoyo incondicional para la culminación de este proyecto.

A mi hijo, Santiago Suárez, quien desde que llego a mi vida es mi mayor inspiración, y con quien hemos demostrado que todo lo podemos lograr; a Juan Suárez, quien me apoya, acompaña y no me ha dejado desfallecer, en este logro.

A la Universidad Nacional de Colombia, departamento de biología y el posgrado en microbiología, por permitirme formarme profesionalmente; a Socorro Prieto, por su apoyo incondicional.

Doy agradecimiento especial a mis tutores, la profesora Nubia Estela Matta y el profesor Mario Vargas-Ramírez, por el apoyo en el desarrollo de esta investigación, por su consejo y ayuda significativa con esta investigación. A la profesora Nubia Estela Matta, un agradecimiento especial por darme la oportunidad de pertenecer al grupo de investigación GERPH.

Agradecimiento especial a todos mis compañeros del grupo de estudio “GERPH: Grupo de estudio de relación parásito–hospedero”, por su apoyo, especialmente a Germán Gutiérrez y Carolina Vargas, por su apoyo y comprensión a lo largo del desarrollo de esta investigación.

A la profesora Martha Calderon, de la Universidad Nacional de Colombia, quien a través de las salidas de campo de la asignatura “Taxonomia animal”, el profesor Oscar Rodríguez de la Fundación Universitaria-Unitrópico y su grupo de investigación, a Rafael Gutiérrez y a Andrés Jiménez, quienes recolectaron algunas de las muestras analizadas; sin sus esfuerzos en campo el desarrollo de esta investigación no hubiera sido posible. VI IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Finalmente, quiero agradecer el apoyo financiero del proyecto Colciencias Número 018- 2018 “FP44842-018-2018” “Hemoparásitos asociados a la herpetofauna colombiana: Aspectos relevantes para su conservación”. Código 111877657872. Resumen y Abstract VII

Resumen

Colombia es un país altamente diverso en flora y fauna, siendo el 4to país con mayor biodiversidad de reptiles en el mundo. Sin embargo, los estudios relacionados con parásitos sanguíneos que infectan animales en vida silvestre son limitados. Apicomplexa es un phylum de protozoarios que agrupa parásitos obligados, como los hemoparásitos que infectan mamíferos, aves y reptiles. Este estudio evaluó la diversidad de hemoparásitos presentes en reptiles de diferentes departamentos, utilizando la determinación morfológica e información del marcador molecular citocromo b, para los géneros Plasmodium y Haemocystidum, y del fragmento de 18S rRNA, para el género Haemogregarinas. Se analizaron 225 individuos, encontrando 148 infectados con Haemogregarinas spp, cuatro con Plasmodium spp., y tres con Haemocystidium spp. Para Plasmodium kentropyxi y Plasmodium carmelinoi, encontrados en Cnemidophorus cf. gramivagus y Ameiva ameiva, respectivamente, se reportó por primera vez linajes de cytb que puden ser usados como BarCode (Capítulo 1). Se reporta la presencia de Haemocystidium sp en Podocnemis vogli (Capítulo 2). Se identificaron 14 secuencias de 18S rRNA asociados a dos morfotipos de Haemogregarinas spp. en Podocnemis vogli, y una secuencia asociada al único morfotipo en Podocnemis unifilis (Capítulo 3). El marcador citocromo b es útil para usarse como BarCode para identificar especies de haemosporidos; en el caso de las Haemogregarinas, la información obtenida con el marcador 18S rRNA, debe complementarse, por lo que sugiere la búsqueda de nuevos marcadores, que contribuyan a mejorar la resolución de las relaciones filogenéticas y la diferenciación entre e intra especies del suborden Adeleorina.

Palabras clave: Tortugas, Apicomplexa, citocromo b, 18S rRNA.

VII IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES I DEPARTAMENTOS DE COLOMBIA.

Abstract

Colombia is a highly diverse country in flora and fauna, being the fourth country with the greatest reptile biodiversity in the world. Nonetheless, the studies related to blood parasites that infect wildlife animals are limited. Apicomplexa is a protozoan’s phylum that groups obligate parasites, comprising hemoparasites infecting mammals, birds, and reptiles. In this study, the diversity of hemoparasites present in reptiles of different departments were evaluated by using morphological determination and molecular information of Cytochrome b molecular marker for Plasmodium and Haemocystidium genera, and a fragment of 18S rRNA for the Haemogregarina genus. 225 individuals were analyzed, 147 of them were positive for Haemogregarinas spp. infection, four for Plasmodium spp. and three for Haemocystidium spp. For Plasmodium kentropyxi y Plasmodium carmelinoi, found in Cnemidophorus cf. gramivagus y Ameiva ameiva, respectively, cytb linages that could be used as BarCode were reported for the first time (Chapter one). The Haemocystidium sp. was reported in Podocnemis vogli (Chapter two). 14 sequences of 18S rRNA were identified and associated with two morphotypes of Haemogregarinas spp. in Podocnemis vogli and one sequence was associated with a unique morphotype in Podocnemis unifilis (Chapter 3). Cytochrome b is a useful molecular marker as BarCode for haemosporidian species identification; the information obtained with 18S rRNA molecular marker was not sufficient for species identification for Haemogregarinas, therefore, we suggest to seek new markers that contribute to the resolution of phylogenetic relationships and the differentiation between and within species of Adelorina suborder.

Keywords: Turtle, tortoise, Apicomplexa, Cytochrome b, 18S rRNA

Contenido IX

Contenido

Lista de figuras ...... XII Lista de tablas ...... XIV Objetivos XV Objetivos específicos ...... XV Introducción ...... 17 Referencias ...... 20 Revisión de literatura ...... 24 Clasificación taxonómica...... 24 Orden Haemosporidia (Haemosporina) ...... 26 Familia Plasmodiidae...... 27 Plasmodium spp...... 27 Familia Garnidae...... 28 Familia Haemoproteidae...... 28 Ciclo de vida...... 29 Análisis filogenético...... 30 Suborden Adeleorina ...... 31 Familia Haemogregarinidae...... 31 Familia Hepatozoidae ...... 31 Familia Karyolysidae: ...... 31 Familia Dactylosomatoidae...... 32 Ciclo de vida ...... 32 Marcadores moleculares usados para en el estudio del suborden Adeleorina en herpetos ...... 33 Referencias ...... 34

1. Capítulo 1. Plasmodium parasites in reptiles from the Colombia Orinoco-Amazon basin: a re-description of Plasmodium kentropyxi Lainson R, Landau I, Paperna I, 2001 and Plasmodium carmelinoi Lainson R, Franco CM, da Matta R, 2010 38 1.1 Abstract ...... 41 1.2 Introduction ...... 41 1.3 Materials and methods ...... 44 1.3.1 Study area ...... 44 X IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

1.3.2 Samples ...... 45 1.3.3 Microscopic analyses ...... 45 1.3.4 Detection of haemosporidian parasites by polymerase chain reaction (PCR) 46 1.3.5 Phylogenetic analysis of the cyt b fragment ...... 46 1.4 Results ...... 47 1.4.1 Molecular phylogenies ...... 58 1.5 Discussion ...... 61 1.6 Acknowledgements ...... 66 1.7 Reference ...... 67 1.8 Appendix ...... 71 2. Capítulo 2. Haemocystidium spp., a species complex infecting ancient aquatic turtles of the family Podocnemididae: First report of these parasites in Podocnemis vogli from the Orinoquia ...... 80 2.1 Abstract ...... 82 2.2 Introduction ...... 82 2.3 Materials and methods ...... 83 2.3.1 Sampling ...... 83 2.3.2 Ethical statement and sampling permits ...... 84 2.3.3 Blood film examination ...... 84 2.3.4 DNA extraction and detection of haemosporidians parasites by a polymerase chain reaction ...... 84 2.3.5 Phylogenetic analysis of the parasite cytb gene fragment ...... 84 2.4 Results ...... 85 2.4.1 Morphological detection of Haemosporidians parasites ...... 85 2.4.2 Genetic distances and phylogenetic relationships of Haemocystidium genus 86 2.4.3 Morphological description ...... 87 2.5 Discussion ...... 87 2.6 Conclusions ...... 90 2.7 Acknowledgements ...... 91 2.8 Reference ...... 91 2.9 Appendix ...... 93

3. Capítulo 3. The puzzle of Haemogregarine infection in aquatic South American turtles...... 97 3.1 Abstract ...... 98 3.2 Introduction ...... 99 3.3 Materials and methods ...... 102 3.3.1 Sampling ...... 102 3.3.2 Morphological and morphometric analysis...... 104 3.3.3 Molecular analysis ...... 104 3.4 Results ...... 106 3.4.1 Morphological and morphometric analyses ...... 106 3.4.2 Genetic distances and phylogenetic relationships ...... 109 3.5 Discussion ...... 115 3.6 Conclusion and perspectives ...... 122 3.7 Acknowledgements ...... 122 3.8 Reference ...... 123 3.9 Appendix ...... 130 Contenido XI

4. Capítulo 4. Discusión general ...... 131 4.1 Referencia ...... 137

5. Conclusiones y recomendaciones ...... 142 5.1 Conclusiones ...... 142 5.2 Recomendaciones ...... 144

A. Anexo: Divulgación...... 146 Presentación Oral...... 146 Posters ...... 148 Libro...... 150 Bibliografía ...... 166

XII IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Lista de figuras Capítulo 1

Figure I 1 Map of Guaviare Department-Colombia, showing sampling locations. Geographic coordinates of the sampling points are in Online Resource 1. Cerro Azul; Ciudad de Piedra; Playa Güio; Puentes Naturales; Puerta de Orión……………………..44

Figure I 2 Plasmodium (Sauramoeba) kentropyxi blood stages infecting Cnemidophorus cf. gramivagus Mccrystal & Dixon, 1987 (a) multiple infection in the same erythrocyte (a-c) Meronts (d) Immature gametocyte (e) Mature macrogametocyte, where a vacuole of <1.1 µm is observed (f) Microgametocyte. Plasmodium (Carinamoeba) carmelinoi blood stages infecting Ameiva ameiva Linnaeus, 1758 (g) Trophozoite (h-j) Meronts (h) tinny meronts white arrow head (j) pigment inside vacuole (black arrow head) (k) Meront with uncommon distribution (l) Gametocyte with a small vacuole (black arrow head). Plasmodium sp., infecting Anolis auratus Daudin, 1802 (m) Double infection in an erythrocyte with trophozoites (n) Non-segmented meront (o) Segmented meront (p) Gametocyte. Plasmodium (Lacertamaoeba) sp. (q) Immature meront (r) Mature meronts (s-t) Gametocytes. Black arrow heads parasite vacuole, and white arrow pigment granules. Giemsa-stained thin blood films. Scale bar=10 μm………………………………………….56

Figure I 3 A Bayesian phylogenetic hypothesis of lizard haemosporidian parasites based on partial sequences of cyt b gene (74 sequences and 500 bp excluding gaps). The values above branches are posterior probabilities (see Material and Methods). Leucocytozoon genus was used as outgroup……………………………………………………………………59

Figure I 4 A Bayesian phylogenetic hypothesis of lizard haemosporidian parasites based on partial sequences of cyt b gene (70 sequences and 981 bp. excluding gaps). The values above branches are posterior probabilities (see Material and Methods). The short sequences of two-lizard parasites P. zonuridae and P. intabazwe and the two sequences of ungulate Plasmodium were excluded from this analysis. Leucocytozoon genus was used as outgroup……………………………………………………………………………………….60 Capítulo 2

Figure II 1 Timeline of the taxonomic classification of the genus Haemocystidium. Relevant events in the identification of gender…………………………………………………… …… 83

Figure II 2 Species and sampling locations for turtle analyzed in the study. Turtle species present in these localities are shown. ………………………………………………… 84 Contenido XIII

Figure II 3 (A) A Bayesian phylogenetic análisis of reptile haemosporidian parasites basad on 62 partial sequences of cytb gene sequences corresponding to a bigger fragment of cytb (707 bp excluding gaps). Leucocytozoon genus was used as out group. In parenthesis are GenBank sequence accession number, isolate name, and turtle species name respectively. Branch color indicates the parasites genus: Blue, Plasmodium sp.; light green, Haemocystidium spp. Infecting lizards and snakes; dark green, Haemocystidium sp. Infecting turtles; black, Haemoproteus and Leucocytozoon spp. (B–C) Estimates of evolutionary divergence between/within Haemocystidium spp.Genetic distances were estimated using the bigger fragment of cytb (707 bp excluding gaps). The number of base substitutions per site between sequences are shown in black and the standard error estimate(s)are shown above the diagonal in blue. Evolutionary divergence between/within Haemocystidium pacayae and Haemocystidium sp. (GERPH:PC005) are shown in bold and red respectively. Sequences previously identiﬁed as Hae.pacayae (KF049495 and KF049507) are likely Hae. (Simondia) sp. (group2). (For interpretation of the references to color in this ﬁgure legend, the reader is referred to the Web version of this article 86

Figure II 4. Haemocystidium (Simondia) sp. (GERPH: PC005) identified in Podocnemis vogli (A–H). (A–F) Young gametocytes, (E) coinfection with a gamonte of Haemogregarina, (G) microgametocyte, and (H) macrogametocyte. Haemocystidium (S.) pacayae (GERPH: PC004-PC006) identified in P. Vogli (I–J). (I) Young gametocyte, (K) microgametocyte, and (L) macrogametocyte. Bold black arrow: hemozoingranules; white arrow: parasitophorous vacuole; fine black arrow: vacuole. Giemsa stain, Scale bar: 10 μm……………………… 86

Capítulo 3

Figure III 1. Species and sampling locations for turtle analyzed in the study. In red localities were P. vogli come from……………………………………………………………………….102

Figure III 2. Haemogregarina sp. identified in Podocnemis vogli and P. unifilis. Haemogregarina sp. In P. vogli (A) Meront (B-I) Morphotype 1. (B-F) Young gamont (G-I) Mature gamont. (J-L) Morphotype 2. Mature gamont. Arrow double: Acytoplasmic space; double triangle: nucleus. Black arrows: cytoplasm pink hue. Haemogregarina sp. In P. unifilis (M-P) Mature gamont. Double triangle: nucleus. Bar 10 μm………………………109

Figure III 3. A Maximum likelihood estimation hypothesis of Haemogregarina parasites based on partial sequences of 18S rRNA gene (108 sequences and 595 bp). Adelina dimidiata genus used as outgroup…………………………………………………………….112

Figure III 4. A Bayesian phylogenetic hypothesis of Haemogregarina parasites based on partial sequences of 18S rRNA gene (108 sequences and 595 bp). The values above branches are posterior probabilities). Adelina dimidiata was used as outgroup………..113 XIV IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Lista de tablas

Capítulo 1

Table I 1. Reptiles sampled in this study and the prevalence of Plasmodium spp. 47

Table I 2 Morphometric characteristics of the Plasmodium species found in the reptiles sampled in this study. Measurements are in micrometers (maximum and minimum values). Average followed by the standard deviation for each morphological characteristic is given. 57 Capítulo 2

Table II 1 Turtle species analyzed in the present study, from diverse localities in Colombia. Common names, size sample, and locality are shown. *The only locality that has turtles infected with Haemocystidium……………………………………………………………… 85

Table II 2. Morphometrical features of Haemocystidium (Simondia) complex species described in South America and their host. Measurements of infected and non-infected erythrocytes are shown……………………………………………………………………… 87

Table II 3. Genetic distances using the cytb or mDNA, reported in haemosporidian species, indicating if they are cryptic species or not………………………………………………….. 90

Capítulo 3

Table III 1. Turtle species analyzed in the present study, from diverse localities in Colombia. 103

Table III 2. Parasitaemia and sequence of each sample analyzed of P. vogli of department Vichada and Casanare 114

Contenido XV

Objetivos Evaluar la diversidad de linajes moleculares, morfotipos y rango de hospederos, relacionados con hemoparásitos que infectan algunas especies de reptiles presentes en diferentes departamentos de Colombia.

Objetivos específicos  Establecer la prevalencia, parasitemia y rango de hospedero de los hemoparásitos presentes en las muestras.  Determinar los morfotipos presentes mediante características morfológicas y morfométricas de los hemoparásitos encontrados.  Determinar los linajes moleculares de cyt b y 18s RNA, de los hemoparásitos encontrados.  Analizar la asociación entre linajes moleculares y morfotipos de los géneros de parásitos encontrados en el estudio.

IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Introducción La ubicación geográfica de Colombia en Suramérica permite que nuestro país sea un puente entre América central y del Sur, lo cual, junto con las diferencias climáticas, hidrográficas y geográficas del territorio, permiten la presencia de diversos ecosistemas, fauna y flora. A nivel mundial, el 70% de la biodiversidad se encuentra albergada en 17 países, dentro de los cuales se encuentra Colombia (Velandia 2019), como consecuencia de la diversidad de climas y hábitats que conforman el territorio.

Colombia ocupa el cuarto lugar a nivel mundial en cuanto a biodiversidad de reptiles se refiere, reportándose cerca de 510 especies distribuidas en el territorio, de las cuales 43 se han clasificado en diversas categorías de amenaza, y 102 especies aun no cuentan con la información suficiente (Moreno et al. 2016), por lo cual, los conocimientos relacionados con su entorno, hábitats y características inherentes a los individuos toman relevancia en la actualidad. Dentro de las amenazas que se han reportado afectando a este grupo de organismos se encuentra: el tráfico ilegal de especies, el consumo de carne o huevos, la pérdida, transformación y degradación del hábitat (Andrade 2011; Moreno et al. 2016;

Ortiz-Moreno and Rodríguez-Pulido 2017), sin embargo, a pesar de la biodiversidad de herpetos en el territorio, el conocimiento relacionado con los organismos que pueden infectarlos como los parásitos aún es escaso.

18 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Dentro de los estudios desarrollados en Colombia, relacionados con microorganismos en herpetos, se ha reportado organismos asociados al caparazón de las tortugas (Donato-

Rondon et al. 2018), la infección de Salmonella sp. en Crocodylus intermedius y Testudinos

(Pachón et al. 2011), de trematodos (Fernandes and Kohn 2014), y parásitos

Monogenoidea (Cohen et al. 2013); además, se ha reportado la presencia de ectoparásitos asociados a tortugas (Garcés-Restrepo et al. 2013) y de Rickettsia spp. en Amblyomma dissimile (Santodomingo et al. 2018). No obstante, a la fecha se desconoce la biodiversidad de hemoparásitos que infectan fauna de vida silvestre, y los efectos que la infección pueda generar.

Los parásitos representan cerca del 50% de las especies existentes en el planeta, por lo cual, poseen un papel relevante en la regulación de los ecosistemas, impactando generalmente de forma negativa las especies a las cuales infecta, relacionándose con eventos de emergencia y reemergencia de enfermedades infecciosas, y considerándose un pool zoonótico, que puede afectar a animales domésticos o ser humano (Daszak 2000).

El phylum Apicomplexa, agrupa a protozoos intracelulares obligados, incluyendo alrededor de 6000 especies a la fecha (Adl et al., 2007), a pesar que se considere que esta cifra corresponde solamente al 0,1% de las especies existentes (Morrison 2009). Dentro de este phylum se encuentran los hemoparásitos que pueden infectar el torrente sanguíneo de mamíferos, aves, y herpetos. Específicamente, en herpetos se ha reportado la presencia de los géneros Plasmodium, Haemocystidium, Garnia, Haemogregarina, Hepatozoon,

Hemolivia, Karyolysus, y Dactylosoma (Barta 1991; Telford 2009; Lainson 2012; Tomé et al. 2014; Cook et al. 2015).

Actualmente, la identificación de los hemoparásitos que infectan reptiles se basa en la caracterización morfológica de los estadios parasitarios identificados en el hospedero vertebrado, la caracterización del ciclo en el vector, y la amplificación de un fragmento de Introducción 19 los genes citocromo b (cytb) o 18S rRNA para Haemosporidios o géneros pertenecientes al suborden Adeleorina, respectivamente; a pesar que ya se encuentra disponible el genoma mitocondrial de Hepatozoon catesbianae en Lithobates clamitans (Leveille et al.

2014), es para la única especies que infecta herpetos, que se cuenta con esta información.

El estudio de los organismos pertenecientes a este grupo a lo largo de la historia se ha centrado en géneros como Plasmodium, teniendo en cuenta la relevancia en salud pública que posee. No obstante, el estudio géneros como Haemogregarina o Hepatozoon han aumentado considerablemente en las últimas décadas (Motta et al. 2011; O’Dwyer et al.

2013; Cook et al. 2014, 2016; Úngari et al. 2018). A nivel mundial, se han desarrollado diversos estudios enfocados en determinar la presencia de estos hemoparásitos, así como, en análisis filogenéticos, caracterización e identificación morfológica y molecular (Siroky et al. 2007; Davis and Sterrett 2011; Perkins 2014; Tomé et al. 2014; Cook et al. 2015; Maia et al. 2016) los cuales, aportan nueva información taxonómica, de rango de hospederos o geográficos.

En Suramérica, el estudio relacionado con este grupo se ha centrado en países como

Brasil (Lainson et al. 2003; Dantas et al. 2008; Lainson 2012; Soares et al. 2014, 2017;

Úngari et al. 2018) o Perú (Paperna et al. 2009; Pineda-Catalan et al. 2013). En Colombia, la caracterización de hemoparásitos en la herpetofauna se limitó hasta hace algunos años a la década de los noventa, en localidades como los Llanos Orientales, Valle del río Cauca,

San Andrés y Providencia, y Antioquia (Ayala et al. 1973; Ayala 1975; Ayala y Spain 1976); no obstante, las investigaciones desarrolladas por nuestro grupo de investigación (Moreno et al. 2015; Gutierrez 2016) demuestran el potencial que tiene este tipo de investigaciones en Colombia. 20 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

El objetivo de este estudio está enfocado en evaluar la diversidad de linajes moleculares, morfotipos y rango de hospederos, relacionados con hemoparásitos que infectan algunas especies de reptiles presentes en diferentes departamentos de Colombia.

Referencias

Adl, S. M., Leander, B. S., Simpson, A. G., Archibald, J. M., Anderson, O. R., Bass, D., ... & Kolisko, M. (2007). Diversity, nomenclature, and taxonomy of protists. Systematic Biology, 56(4), 684-689.

Andrade C MG (2011) Estado del conocimiento de la biodiversidad en Colombia y sus amenazas. Consideraciones para fortalecer la interacción ambiente-política. Rev Acad Colomb Cienc 35:491–507

Ayala SC (1975) Malaria and hemogregarines from lizards of the Western Caribbean Islands of San Andrés and Providencia. Rev Inst Med Trop Sao Paulo 17:218–224

Ayala S.C, D’Alessandro A, Mackenzie R., Angel D (1973) Hemoparasite Infections in 830 Wild Animals from the Eastern Llanos of Colombia. The Journal of Parasitology 59:52–59. doi: 10.2307/3278571

Ayala S.C, Spain JL (1976) A population of Plasmodium colombiense sp. n. in the iguanid lizard, Anolis Auratus. J Parasitol 62:177–189

Barta JR (1991) The Dactylosomatidae. In: Advances in Parasitology. Elsevier, pp 1–37

Cohen SC, Justo MCN, Kohn A (2013) South American Monogenoidea parasites of fishes, amphibians and reptiles. South American Monogenoidea parasites of fishes, amphibians and reptiles

Cook CA, Lawton SP, Davies AJ, Smit NJ (2014) Reassignment of the land tortoise haemogregarine Haemogregarina fitzsimonsi Dias 1953 (Adeleorina: Haemogregarinidae) to the genus Hepatozoon Miller 1908 (Adeleorina: Hepatozoidae) based on parasite morphology, life cycle and phylogenetic analysis of 18S rDNA sequence fragments. Parasitology 141:1611–1620. doi: 10.1017/S003118201400081X

Cook CA, Netherlands EC, Smit NJ (2015) First Hemolivia from southern Africa: reassigning chelonian Haemogregarina parvula Dias, 1953 (Adeleorina: Haemogregarinidae) to Hemolivia (Adeleorina: Karyolysidae). African Zoology 50:165–173. doi: 10.1080/15627020.2015.1044467

Cook CA, Netherlands EC, Smit NJ (2016) Redescription, molecular characterisation and taxonomic re-evaluation of a unique African monitor lizard haemogregarine Introducción 21

Karyolysus paradoxa (Dias, 1954) n. comb. (Karyolysidae). Parasites & Vectors 9:. doi: 10.1186/s13071-016-1600-8

Dantas FT, Oliveira EFF, Soares FÂM, et al (2008) Ticks infesting amphibians and reptiles in Pernambuco, Northeastern Brazil. Revista Brasileira de Parasitologia Veterinária 17:218–221. doi: 10.1590/S1984-29612008000400009

Daszak P (2000) Emerging Infectious Diseases of Wildlife-- Threats to Biodiversity and Human Health. Science 287:443–449. doi: 10.1126/science.287.5452.443

Donato-Rondon JCh, González-Trujillo JD, Romero B, Castro-Rebolledo MI (2018) Diatom assemblages associated with turtle carapaces in the Neotropical region. RBT 66:1362. doi: 10.15517/rbt.v66i4.31396

Fernandes BMM, Kohn A (2014) South American trematodes parasites of amphibians and reptiles. South American trematodes parasites of amphibians and reptiles

Garcés-Restrepo MF, Giraldo A, Carr JL, Brown LD (2013) Turtle ectoparasites from the Pacific coastal region of Colombia. Biota Neotrop 13:74–79. doi: 10.1590/S1676- 06032013000300009

Gutierrez G (2016) PARÁSITOS SANGUÍNEOS EN ALGUNOS REPTILES DE LA LOCALIDAD DE YONDÓ, ANTIOQUIA, COLOMBIA. Universidad Nacional de Colombia

Lainson R (2012) Atlas of protozoan parasites of the amazonian fauna of Brazil. Vol. 1. Haemosporida of reptiles.

Lainson R, Souza MC de, Franco CM (2003) Haematozoan parasites of the lizard Ameiva ameiva (Teiidae) from Amazonian Brazil: a preliminary note. Memórias do Instituto Oswaldo Cruz 98:1067–1070. doi: 10.1590/S0074-02762003000800016

Leveille AN, Ogedengbe ME, Hafeez MA, et al (2014) The Complete Mitochondrial Genome Sequence of Hepatozoon catesbianae (Apicomplexa: Coccidia: Adeleorina), a Blood Parasite of the Green Frog, Lithobates (Formerly Rana clamitans). Journal of Parasitology 100:651–656. doi: 10.1645/13-449.1

Maia, D. JamesHarris, Salvador Carranza (2016) Reconstruction of the evolutionary history of Haemosporida (Apicomplexa) based on the cyt b gene with characterization of Haemocystidium in geckos (Squamata: Gekkota) from Oman. Parasitology international, 65:5–11

Moreno A, González L, Barreto D (2015) CARACTERIZACIÓN MORFOLÓGICA DE HEMOPARÁSITOS PRESENTES EN ALGUNOS REPTILES Y ANFIBIOS DE GUAVIARE, COLOMBIA. Universidad Colegio Mayor de Cundinamarca 22 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Moreno LA, Andradre GI, Ruíz-Contreras LF, (Eds.) (2016) Biodiversidad 2016. Estado y tendencia de la biodiversidad de Colombia.

Morrison DA (2009) Evolution of the Apicomplexa: where are we now? Trends in Parasitology 25:375–382. doi: 10.1016/j.pt.2009.05.010

Motta ROC, Cunha LM, Leite RC, et al (2011) Hepatozoon spp. (Apicomplexa: Hepatozoidae) infection and selected hematological values of the neotropical rattlesnake, Crotalus durissus collilineatus (Linnaeus, 1758) (Serpentes: Viperidae), from Brazil. J Zoo Wildl Med 42:399–407

O’Dwyer LH, Moço TC, Paduan K dos S, et al (2013) Description of three new species of Hepatozoon (Apicomplexa, Hepatozoidae) from Rattlesnakes (Crotalus durissus terrificus) based on molecular, morphometric and morphologic characters. Experimental Parasitology 135:200–207. doi: 10.1016/j.exppara.2013.06.019

Ortiz-Moreno ML, Rodríguez-Pulido JA (2017) Estado del conocimiento y amenazas de la tortuga sabanera (Podocnemis vogli, Podocnemididae) en Colombia. Orinoquia 21:26. doi: 10.22579/20112629.391

Paperna I, Bastien P, Chavatte J-M, Landau I (2009) Lankesterella poeppigii n. sp. (Apicomplexa, Lankesterellidae) de Bufo poeppigii (Tschudi, 1845) del Perú. Revista peruana de biología 16:165–168

Perkins SL (2014) Malaria’s Many Mates: Past, Present, and Future of the Systematics of the Order Haemosporida. Journal of Parasitology 100:11–25. doi: 10.1645/13-362.1

Pineda-Catalan O, Perkins SL, Peirce MA, et al (2013) Revision of Hemoproteid Genera and Description and Redescription of Two Species of Chelonian Hemoproteid Parasites. Journal of Parasitology 99:1089–1098. doi: 10.1645/13-296.1

Santodomingo A, Cotes-Perdomo A, Foley J, Castro LR (2018) Rickettsial infection in ticks (Acari: Ixodidae) from reptiles in the Colombian Caribbean. Ticks and Tick-borne Diseases 9:623–628. doi: 10.1016/j.ttbdis.2018.02.003

Siroky P, Kamler M, Frye FL, et al (2007) Endogenous development of Hemolivia mauritanica (Apicomplexa: Adeleina: Haemogregarinidae) in the marginated tortoise Testudo marginata (Reptilia: Testudinidae): evidence from experimental infection. Folia Parasitologica 54:13–18. doi: 10.14411/fp.2007.002

Soares HS, Marcili A, Barbieri ARM, et al (2017) Novel piroplasmid and Hepatozoon organisms infecting the wildlife of two regions of the Brazilian Amazon. International Journal for Parasitology: Parasites and Wildlife 6:115–121. doi: 10.1016/j.ijppaw.2017.05.002 Introducción 23

Soares P, de Brito, E. S, Paiva, F., Pavan, D, Viana, L. A. (2014) Haemogregarina spp. in a wild population from Podocnemis unifilis Troschel, 1848 in the Brazilian Amazonia. Parasitology research 113:4499–4503

Telford SRJr (2009) Hemoparasites of the Reptilia: color atlas and text. Taylor & Francis, Boca Raton

Tomé B, Maia JP, Salvi D, et al (2014) Patterns of genetic diversity in Hepatozoon spp. infecting snakes from North Africa and the Mediterranean Basin. Systematic Parasitology 87:249–258. doi: 10.1007/s11230-014-9477-4

Úngari LP, Santos ALQ, O’Dwyer LH, et al (2018) Haemogregarina podocnemis sp. nov.: description of a new species of Haemogregarina Danilewsky 1885 (Adeleina: Haemogregarinaidae) in free-living and captive yellow-spotted river turtles Podocnemis unifilis (Testudines: Podocnemididae) from Brazil. Parasitology Research 117:1535–1548. doi: 10.1007/s00436-018-5817-7

Velandia O (2019) La biodiversidad y los servicios ecosistémicos. http://www.humboldt.org.co/es/biodiversidad/que-es-la-biodiversidad. Accessed 2 Mar 2019

24 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Revisión de literatura El phylum Apicomplexa comprende parásitos intracelulares obligados que infectan una gran variedad de animales, caracterizados por la presencia de un complejo apical usado para invadir células hospederas (Votýpka et al. 2017). Dentro de este phylum se incluye patógenos de importancia para la salud pública como Plasmodium o Toxoplasma y otros considerados parásitos emergentes como Babesia y Cryptosporidium. En cuanto al origen de estos parásitos, se ha sugerido que inicialmente invadieron invertebrados marinos, análisis moleculares han permitido ubicar su origen entre 600-800 millones de años antes de la emergencia de los vertebrados.

Clasificación taxonómica La clasificación taxonómica de los parásitos de interés en el presente documento, inicialmente se basó en característica morfológicas, sin embargo, esta clasificación ha cambiado, como consecuencia de la utilización de marcadores moleculares, específicamente del 18S rRNA, único marcador utilizado en este grupo para la realización de análisis filogenéticos y los nuevos taxones muestreados. Acorde con los grupos de interés que abordará este estudio, este documento seguirá la taxonomía propuesta por Adl et al., (2019). De acuerdo con esta clasificación, dentro del phylum Apicomplexa, se encuentra el Subphylum: Conoidasida, Clase coccidia, Suborden Adeleorina: con las familias Haemogregarinidae; Hepatozoidae, Dactylosomatidae, Karyolysidae, y Adeleidae, Klosiellidae y Legerellidae, que corresponden a parásitos con ciclos de vida monoxenos que infectan invertebrados marinos (Tabla 1 y 2). El suborden Adeleorina, se caracteriza por el desarrollo sexual mediante “sicigia”, proceso que está ausente en el suborden Eimeriorina (Adl et al. 2012).

Introducción 25

Tabla 1. Clasificación de Apicomplexa (Levine 1970) Tomado y adaptado de Votýpka et al (2017).

Parásitos obligados Subphylum: Conoidasida Levine, 1988 Clase: Gregarinida Dufour, 1828 Gregarines Archigregarinorida Grasse 1953 Eugregrarinorida Léger 1900 Clase: Coccidia Leuckart, 1879 Coccidia, haemogregarinas Adeleorina Léger 1911 Eimerionirina Léger 1911 Cryptosporidida incertae sedis Cryptosporidia Subphylum: Aconoidasida Mehlhorn, Peters and Hematozoa Haberkorn, 1980 Clase: Haemosporidia Danielewsky, 1885 Haemosporidia Clase: Piroplasmida Wenyon, 1926 Piroplasmas Clase: Nephromycida Cavalier-Smith

1993, emend. (Adl et al. 2019) Relicto aplicomplexa (vida-libre) Colpodellida Cavalier-Smith, 1993 incertae sedis Colpodellidos Chromerida Moore et al., (2008) incertae sedis Chromeridos

Tabla 2. Detalle de la Clasificación de Apicomplexa Levine (1970) y Adl et al.(2012, 2019), Los géneros indicados con * hacen referencia a aquellos de interés en el proyecto.

Phylum Apicomplexa Clase Conoidasida Subclase Gregarinasina Subclase Coccidia Orden Eucoccidiorida Suborden Eimeriorina Familia Sarcocystidae Toxoplasma Neospora Sarcocystis Familia Eimeridae Eimeria Isospora Caryosporoa Familia Lankestellidae Lainsonia Schellackia* Lankesterella Orden Eucoccidiorida Suborden Adeleorina Familia Haemogregarinidae Haemogregarina* Desseria 26 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Cyrilia Familia Hepatozoidae Hepatozoon* Familia Karyolysidae Hemolivia* Karyolysus* Familia Dactylosomatidae Babesiosoma Dactylosoma* Familia Legerellidae Familia Klosiellidae Familia Adeleidae Clase Agamococcidiorida Clase Protococcidiorida Clase Aconoidasida Orden Haemosporidia (Haemosporina) Familia Haemoproteidae Género Haemoproteus (aves) Subgénero Haemoproteus Subgénero Parahaemoproteus Género Haemocystidium* Subgénero Simondia Subgénero Haemocystidium Familia Plasmodidae Género Plasmodium* Subgénero Sauramoeba Subgénero Carinamoeba Subgénero Lacertamoeba Subgénero Paraplasmodium Subgénero Ophidiella Género Hepatocystis Género Polychromophilus Familia Leucocytozoidae Género Akiba Género Leucocytozoon Familia Garniidae Género Garnia* Género Fallisia

Orden Haemosporidia (Haemosporina)

El orden Haemosporidia pertenece a la clase Aconoidasida, la cual se caracteriza por la ausencia del conoide en la estructura del complejo apical. Los protozoarios pertenecientes a este orden son transmitidos por diversorios vectores (artrópodos hematófagos), dentro de este orden se encuentra el género Plasmodium, agente causal de la malaria humana. La mayoría de las especies descritas en este orden se han descrito infectando aves y Introducción 27 reptiles.

Este orden está compuesto por cuatro familias, las cuales pueden diferenciarse morfológicamente en el hospedero vertebrado: Plasmodiidae (posee gránulos de hemozoina y merogonia eritrocitaria); Haemoprotidae (posee gránulos de hemozoína, ausencia merogonia eritrocitaria); Garniidae (ausencia de gránulos de hemozoína y presencia de merogonia eritrocitaria) y Leucocozytoidae (Ausencia de gránulos de hemozoína y de merogonia eritrocitaria) (Perkins 2014).

Familia Plasmodiidae. Esta familia agrupa tres géneros Hepatocystis, que infecta mamíferos, Polychromophilus, que infecta murciélagos y Plasmodium, que ha sido reportado infectando mamíferos, aves y reptiles.

Plasmodium spp.

Este género agrupa los agentes de la malaria humana, por lo cual, ha sido el género mejor caracterizado de este orden. A nivel taxonómico se han reportado más de 200 especies, de las cuales se estima que cerca de 100 especies infectan reptiles. La descripción de especies que infectan estos organismos, inicialmente se basó en caracteres morfológicos (Garnham 1966; Telford 1988); en reptiles, por ejemplo, el tamaño de los merontes eritrocitarios. Con base en esta característica Garnham (1966) propuso la implementación de 3 subgéneros, para las especies que infectan lagartos Sauramoeba (Merontes grandes), Carinamoeba (merontes pequeños) y Ophidiella, el cual agrupa las especies de Plasmodium que infectan serpientes. Posteriormente, Telford (1988), propuso 4 subgéneros más: Paraplasmodium, Asiamoeba, Lacertamoeba y Garnia (Tabla 3).

28 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Tabla 3. Clasificación de subgéneros del género Plasmodium, con base en características morfológicas de gametocitos y merontes

Característica Merontes Subgénero Gametocitos Divisiones Tamaño nucleares Sauramoeba Grandes Grande 4-7 Carinamoeba Pequeño Pequeño 2-3 Lacertamoeba Mediano Mediano 3 -5 Paraplasmodium Grandes Mediano --- 4 a 15 veces más grande que Asiamoeba ------esquizontes

En relación con el subgénero Garnia, la clasificación taxonómica a la fecha continúa siendo controversial; algunos autores consideran a Garnia como una familia aparte.

Familia Garnidae.

Esta familia, agrupa tres géneros Garnia, Fallisia y Progarnia. A diferencia de los géneros Plasmodium y Haemoproteus, estos géneros se caracterizan por la presencia de merogonia eritrocitaria y la ausencia de gránulos de hemozoina. Para el género Garnia, reportado en aves y reptiles, se han descrito 11 especies infectando lagartos del Neotrópico (Brasil, Haití, Panamá y Venezuela); Fallisia se ha reportado infectando solamente trombocitos o leucocitos (Lainson et al. 1971, 1974; Valkiūnas 2005), y Progarnia, que se ha reportado infectando cocodrilos.

Familia Haemoproteidae. La clasificación taxonómica de los dos géneros: Haemoproteus (infecta aves) y Haemocystidium (infecta reptiles), que conforman esta familia ha sido controversial a lo largo de la historia. Haemoproteus, fue identificado en aves por Kruse en 1890, quien creó este género; mientras que Haemocystidium, fue creado por Castellani y Willey (1904), infectando reptiles; no obstante, de 1915 a 1966 el género Haemocystidium, fue sinonimizado con Plasmodium (Wenyon 1915) y Haemoproteus (Wenyon 1926; Levine, 1988). Actualmente, Haemoproteus y Haemocystidium se consideran como géneros diferentes, con base en la ausencia de formación de “pseudocytometeros” en los merontes (ubicados en los órganos) de Haemocystidium a diferencia de los de Haemoproteus Introducción 29

(Telford 1996). Mediante análisis moleculares, Pineda- Catalán et al., (2013) demostraron que los haemosporidios que infectan reptiles pertenecen al género Haemocystidium, agrupando en un clado diferente de Haemoproteus, que infectan aves; estos autores, sugieren la implementación de dos subgéneros Haemocystidium (Haemocystidium) que infecta hospederos vertebrados del orden squamatas, desconociéndose el vector y Haemocystidium (Simondia) transmitido por moscas de la familia Tabanidae, infectando quelonios (Telford 1996; Perkins 2014; Maia 2015).

Ciclo de vida.

El ciclo de transmisión de los parásitos mencionados previamente es heteroxeno, involucrando un hospedero invertebrado (vector) y un hospedero vertebrado. A través de la ingesta de formas infectivas presentes en el torrente sanguíneo del hospedero vertebrado por parte del vector, se da paso a la fase merogónica. En este proceso se liberan macro y microgametocitos, los cuales se transforman en macro y microgametos. Los microgametos exflagelan y fertilizan los macrogametos (fecundación), formándose el cigoto, que se transformara en ooquineto. Este, se dirige a las células del intestino delgado del vector, donde se enquista y origina el ooquiste, el cual, sufre esporogonia asexual, generando esporozoítos, que se liberan al hemocele, penetran las glándulas salivales, y en el momento que el vector vuelva a alimentarse de otro hospedero vertebrado, transmitirá la infección (Valkiūnas 2005; Telford 2009). A excepción del cigoto, las demás etapas del ciclo de vida de estos parásitos son haploide, considerándose el ooquineto como un cigoto móvil.

Entre el ciclo de vida de Plasmodium y Haemoproteus existen varias diferencias; en Haemoproteus, los esporozoítos desarrollan tropismo por las células endoteliales, lugar donde se desarrolla la merogonia, liberando cientos o miles de merozoítos. Los merozoítos pueden (1) infectar células del bazo o músculo esquelético (en algunas especies) donde se vuelve a desarrollar la merogonia, y (2) infectar los eritrocitos, donde se desarrollan los gametocitos. En Plasmodium, en el hospedero vertebrado, algunos de los esporozoítos inoculados por el vector se inactivan en las células parenquimatosas, dando como resultados hipnozoítos (Telford 2009). En menor cantidad, otros esperozoítos experimentan el proceso de merogonia, convirtiéndose en criptozoítos, los cuales, pueden infectar macrófagos (generando metacriptozoítos), además, se generan fanerozoítos. 30 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Después de este proceso, los merozoítos generados, se dirigen al endotelio capilar y tejido conectivo de varios órganos e inician más ciclos merogónicos. En los eritrocitos, se desarrollan trofozoítos, merontes micro y macrogametocitos, estadios parasitarios infectivos para el vector (Telford 2009). En el caso del género Haemocystidium, al igual que en el género Haemoproteus, hay ausencia de merogonia eritrocitaria; los merontes de especies como H. papernai se han encontrado en el tejido conectivo y el endotelio de los vasos sanguíneos del corazón, el pulmón, el hígado, riñon y masas musculares estriadas del fémur (Telford 1996).

En reptiles, los estudios relacionados con vectores son limitados, sin embargo, Ayala (1970) describió la fase esporogónica, de Plasmodium mexicanum en Lutzomyia vexator y L. stewarti; en Haemocystidium se ha asociado a Chrysops callidus (Diptera: Tabanidae), como vector de Hae. metchnikovi (DeGiusti et al. 1973)

Análisis filogenético.

La clasificación taxonómica de los Haemosporidia tradicionalmente se basó en las características morfológicas de las formas parasitarias en los hospederos vertebrados, su ciclo de vida, y los vectores que los transmiten. No obstante, esta información resulta insuficiente en el caso de especies crípticas. La implementación de herramientas moleculares ha permitido complementar esta información. Para los Haemosporidios que infectan aves y reptiles, la mayoría de los análisis filogenéticos han sido reconstruidos con base en la información de genes mitocondriales, específicamente del citocromo b. El genoma mitocondrial de estos organismos es una molécula lineal dispuesta en tándem, siendo a la fecha, el más pequeño de los genomas mitocondriales reportados en eucariotas, con un tamaño de 6 kb (Pacheco et al. 2018).

En reptiles, se han descrito alrededor de 100 especies de Plasmodium, con base en información de este marcador. En cuanto a la clasificación taxonómica, el análisis de este marcador, corroboró la clasificación del género Haemoproteus, para los Haemoproteidos que infectan aves y del género Haemocystidium, para aquellos que infectan reptiles, aportando información que soporta la división en dos subgéneros, Haemocystidium y Simondia, los cuales, agrupan como un clado hermano de Plasmodium (Pineda-Catalan et al. 2013). Introducción 31

Suborden Adeleorina

Familia Haemogregarinidae. La familia Haemogregarinidae agrupa tres géneros: Deisseria y Cyrilia (infectan peces) y Haemogregarina, estos últimos son los hemoparásitos mas comunes en tortugas. El género Haemogregarina, se caracteriza por la presencia de gamontes (sin dimorfismo sexual) y merontes en la sangre periférica de los hospederos vertebrados infectados, característica que lo diferencia de géneros como Hepatozoon. Las sanguijuelas son los vectores asociados a su transmisión (Siddall y Burreson 1994).

Familia Hepatozoidae.

El único representante de esta familia, es el género Hepatozoon el cual, es el más comúnmente reportado infectando serpientes, a pesar de encontrarse infectando otros reptiles, aves y mamíferos (Craig et al. 1978; Smith 1996; Merino y Martínez 2014). En la sangre periférica de los hospederos vertebrados, solamente se observan gamontes, los cuales no poseen dimorfismo sexual.

Familia Karyolysidae. Agrupa los géneros Karyolysus y Hemolivia; el género Karyolysus, se ha reportado infectando lagartos y anfibios; los vectores asociados a su trasmisión son los ácaros (Mesosigmata) (Telford 2009; Haklová y Majláthová 2014). Morfológicamente, los gamontes observados en la sangre periférica del hospedero vertebrado, se diferencia de los gamontes de Hepatozoon o Haemogregarina, por lisar el núcleo de la célula hospedera (Telford 2009), efecto del cual se deriva su nombre.

El género Hemolivia es transmitido por garrapatas a tortugas (Telford 2009; Netherlands et al. 2015), a nivel morfológico, en sangre periférica del hospedero vertebrado, los gamontes infectan eritrocitos, poseen forma elíptica o cilíndrica y una cápsula resistente a la tinción; los gamontes maduros poseen el núcleo en posición polar (Siroky et al. 2007). Este género, se diferencia de otros géneros como Hepatozoon por características relacionadas con su ciclo de vida: La esporogonia se divide en dos fases: (1) formación del ooquiste y esporoquinetos; continuando con (2) la formación de esporoquistes y esporozoítos; la merogonia puede ser extra e intra eritrocitaria (Petit et al. 1990). 32 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Familia Dactylosomatoidae.

Agrupa los géneros Babesiosoma y Dactylosoma. Dentro de las características de estos géneros se encuentran, el tamaño pequeño de los gamontes y la ausencia de pigmentos en los gamontes y merogonia eritrocitaria. Dactylosoma produce de 4 a 16 merozoítos, en forma de abanico y Babesiosoma produce menos de 4 merozoítos en forma de roseta o cruz (Hoffman 1999).

Ciclo de vida En general la transmisión de los hemoparásitos del suborden Adeleorina mencionados, se caracterizan por ciclo de vida heteroxenos, donde se involucra diversos hospederos y vectores, los cuales varían de acuerdo con el género analizado. En el caso de Hepatozoon sp., la trasmisión puede darse mediante (1) la ingesta del hospedero invertebrado (sanguijuelas, garrapatas, mosquitos, entre otros) (Smith 1996; Davies and Johnston 2000), (2), por parte del hospedero vertebrado; (2) la ingesta de un hospedero vertebrado mediante el sistema presa-predador (Allen et al. 2011; Tomé et al. 2012) (3) la transmisión a través de la picadura del vector (Desser et al. 1990; Telford 2009) y (4) la transmisión vertical, la cual se ha demostrado para Hepatozoon canis (Murata et al. 1993) y en serpientes vivíparas (Kauffman et al. 2017).

Una vez el hospedero invertebrado (vector) consume sangre infectada con gamontes intraeritrocitarios, estos se dirigen hasta la pared intestinal, lugar donde se desarrolla la sicigia (alineamiento de gametos), la gametogénesis y la fertilización, formando un cigoto. Posteriormente, ocurre la esporogonia, se genera un ooquiste temprano, que se desarrollará en un ooquiste multiesporocístico; este contiene esporozoítos infectivos (ausentes en Haemogregarinas). Los esporozoítos se desarrollan en el hemocele y los túbulos de Malpighi durante la esporogonia (Maia 2015).

Cuando el hospedero vertebrado ingiere el vector infectado, los esporozoítos ingresan al torrente sanguíneo, migran a hígado o pulmones. En estos órganos se forman esquizontes. Se da lugar a la merogonia, los merontes forman quistes, los cuales dan lugar a los merozoítos. Algunos merozoítos y merontes, infectan nuevamente tejido continuando la infección (Telford 2009). Finalmente, los eritrocitos son infectados por merozoítos, se desarrollan trofozoítos, los cuales dan lugar a gamontes maduros mediante la Introducción 33 gamontogonia (Maia 2015).

En relación con su ciclo de vida, dentro de las características que permiten diferenciar los géneros pertenecientes al suborden Adeleorina, en el género Karyolysus, los esporoquistes se desarrollan en el ácaro adulto, y se transmiten transovarcularmente a través del huevo de próxima generación de ácaros, donde se desarrollarán los esporozoítos (Telford 2009); en Dactylosoma, existen diferentes tipos de esquizogonia, una exclusivamente multiplicativa, y otra enfocada en la producción de micro y macrogametocitos.

Marcadores moleculares usados para en el estudio del suborden Adeleorina en herpetos El diagnóstico de las especies pertenecientes a este suborden debe realizarse de manera integral, teniendo en cuenta (1) el desarrollo del parásito en el vector, (2) las características morfológicas de las formas infectivas en el hospedero vertebrado, y (3) la información molecular del mismo.

El marcador molecular utilizado a la fecha para la identificación de estos parásitos es un fragmento del gen nuclear 18S rRNA. En los eucariotas, los genes que codifican rRNA, están organizados en unidades transcriptacionales con un gen de rRNA de subunidad pequeña y otro gen de rRNA de la subunidad grande, los cuales están separados por espaciadores transcritos internos. Estas unidades, se encuentran agrupadas en el genoma, la cual posee múltiples copias repetidas en tándem; en algunos Apicomplexas, como Plasmodium o Theileria, se ha reportado que a pesar que se conserva la estructura de las unidades de DNAr, estas se encuentran dispersas por todo el genoma, así mismo, el número de copias en estos dos organismos, es menor en comparación con organismos como Toxoplasma gondii (110 copias) (Guay, et al., 1992), en Plasmodium se han reportado entre cuatro y ocho copias, y en Theileria tres copias (McCutchan et al, 1995; Kibe et al., 1994) . En Plasmodium, además se han identificado diferentes tipos de rRNA, los cuales se expresan de acuerdo a la etapa del ciclo de vida en la cual se encuentre (Mercereau-Puijalon et al., 2002). En cuanto a la tasa de evolución de estos genes se ha reportado que poseen regiones con diferentes tasas de evolución, específicamente, el gen 18S rRNA, muestra una tasa de evolución lenta, con regiones altamente conservadas (Morand et al. 2015) y otras variables, las cuales son utilizadas para la diferenciación de 34 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA. genéros, en este suborden.

El análisis filogenético del fragmento obtenido para este marcador, a futuro puede variar de acuerdo con las secuencias incluidas, teniendo en cuenta que para géneros como Desseria y Cyrilia aún no se cuenta con información genética asociada; así mismo, la obtención de información genética para especies descritas previamente con base en las características morfológicas ha contribuido a la reasignación de especies entre géneros. En la actualidad, el género Hemolivia agrupa como clado hermano del género Hepatozoon; el género Karyolysus agrupa como basal de los dos géneros mencionados previamente. El género Haemogregarina se ubica como un grupo diferente, en el cual el género basal es Dactylosoma.

La utilización de otros marcadores mitocondriales, de apicoplasto o nucleares, contribuirá a la resolución de las relaciones filogenéticas, y la descripción de especies que puedan identificarse, teniendo en cuenta que a la fecha solamente Hepatozoon catesbianae, posee información del genoma mitocondrial (Leveille et al., 2014).

Referencias Adl SM, Bass D, Lane CE, et al (2019) Revisions to the Classification, Nomenclature, and Diversity of Eukaryotes. Journal of Eukaryotic Microbiology. doi: 10.1111/jeu.12691

Adl SM, Simpson AGB, Lane CE, et al (2012) The Revised Classification of Eukaryotes. Journal of Eukaryotic Microbiology 59:429–514. doi: 10.1111/j.1550-7408.2012. 00644.x

Allen KE, Yabsley MJ, Johnson EM, et al (2011) Novel Hepatozoon in VertebratesFrom the Southern United States. Journal of Parasitology 97:648–653. doi: 10.1645/GE-2672.1

Ayala S.C (1970) Plasmodium mexicanum in California: natural history and development in Phlebotomine sandflies (Diptera: Psychodidae). J Parasitol 56:13

Castellani, A, Willey A (1904) Observations on the haematozoa of vertebrates in Ceylon. Spolia Zeylonic 2:78–92

Craig TM, Smallwood JE, Knauer KW, McGrath JP (1978) Hepatozoon canis infection in dogs: clinical, radiographic, and hematologic findings. J Am Vet Med Assoc 173:967– 972

Chelonian Haemoproteid Haemoproteus metchnikovi by a Tabanid Fly Chrysops callidus. Nature 242, 50–51. https://doi.org/10.1038/242050a0

Desser SS, Siddal ME, Batra JR (1990) Ultrastructural observations on the developmental stages of Lankesterella minima (Apicomplexa) in experimentally infected Rana catesbeiana tadpoles. Journal of Parasitology 76:997–103

Garnham PCC (1966) Malaria Parasites and other Haemosporidia. Blackwell Scientific Publications, Oxford, U. K

Guay J.-M, Huot A., Gagnon S, Tremblay A., Levesque R.C. Physical and genetic mapping of cloned ribosomal DNA from Toxoplasma gondii: primary and secondary structure of the 5S gene Gene, 114 (1992), pp. 165-171

Haklová B, Majláthová V (2014) Phylogenetic relationship of Hepatozoon blood parasites found in snakes from Africa, America and Asia. Parasitology 141:389–398. doi: 10.1017/S0031182013001765

Hoffman (1999) Parasites of North American freshwater fishes. University of California Press, Berkeley, Los Angeles, London

Kauffman KL, Sparkman A, Bronikowski AM, Palacios MG (2017) Vertical Transmission of Hepatozoon in the Garter Snake Thamnophis elegans. Journal of Wildlife Diseases 53:121–125. doi: 10.7589/2016-03-056

Lainson R, Landau I, Shaw JJ (1974) Observations on non-pigmented haemosporidia of Brazilian lizards, including a new species of Saurocytozoon in Mabuya mabouya (Scincidae). Parasitology 69:215–223

Lainson R, Landau I, Shaw JJ (1971) On a new family of non-pigmented parasites in the blood of reptiles: Garniidae fam. nov., (Coccidiida: Haemosporidiidea). Some species of the new genus Garnia . International Journal for Parasitology 1:241–250. doi: 10.1016/0020-7519(71)90027-0

Le Blancq, S. M., Khramtsov, N. V., Zamani, F., Upton, S. J., & Wu, T. W. (1997). Ribosomal RNA gene organization in Cryptosporidium parvum. Molecular and biochemical parasitology, 90(2), 463-478.

Leveille, A. N., Ogedengbe, M. E., Hafeez, M. A., Tu, H. H. A., & Barta, J. R. (2014). The complete mitochondrial genome sequence of Hepatozoon catesbianae (Apicomplexa: Coccidia: Adeleorina), a blood parasite of the green frog, Lithobates (formerly Rana) clamitans. Journal of Parasitology, 100(5), 651-656.

Levine ND (1970) Taxonomy of the sporozoa. J Parasitol 56:208–209 36 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Levine, ND (1988) The protozoan phylum Apicomplex. p. 118-133. 1988. CRC Press, INC, Boca Raton, Florida

Maia JP (2015) Diversity, infection patterns and hostparasite associations of apicomplexan parasites in reptiles. (Doctoral dissertation, Ph. D. Thesis. Universidade do Porto, Porto, Portugal).

McCutchan, T. F., Li, J., McConkey, G. A., Rogers, M. J., & Waters, A. P. (1995). The cytoplasmic ribosomal RNAs of Plasmodium Spp. Parasitology Today, 11(4), 134-138.

Mercereau-Puijalon, O., Barale, J. C., & Bischoff, E. (2002). Three multigene families in Plasmodium parasites: facts and questions. International journal for parasitology, 32(11), 1323-1344.

Merino, Martínez J (2014) First Molecular Characterization of a Hepatozoon Species (Apicomplexa: Hepatozoidae) Infecting Birds and Description of a New Species Infecting Storm Petrels (Aves: Hydrobatidae). The Journal of parasitology 100: doi: 10.1645/13-325.1

Moore, R. B, Oborník, M, Janouškovec, J, et al (2008) A photosynthetic alveolate closely related to apicomplexan parasites. Nature 451:959

Morand S, Krasnov BR, Littlewood DTJ (eds) (2015) Parasite Diversity and Diversification: Evolutionary Ecology Meets Phylogenetics. Cambridge University Press, Cambridge

Murata T, Inoue M, Tateyama S, et al (1993) Vertical transmission of Hepatozoon canis in dogs. J Vet Med Sci 55:867–868

Netherlands EC, Cook CA, Kruger DJD, et al (2015) Biodiversity of frog haemoparasites from sub-tropical northern KwaZulu-Natal, South Africa. International Journal for Parasitology: Parasites and Wildlife 4:135–141. doi: 10.1016/j.ijppaw.2015.01.003

Pacheco MA, Matta NE, Valkiunas G, et al (2018) Mode and Rate of Evolution of Haemosporidian Mitochondrial Genomes: Timing the Radiation of Avian Parasites. Mol Biol Evol 35:383–403. doi: 10.1093/molbev/msx285

Perkins SL (2014) Malaria’s Many Mates: Past, Present, and Future of the Systematics of the Order Haemosporida. Journal of Parasitology 100:11–25. doi: 10.1645/13-362.1

Petit G, Landau I, Baccam D, Lainson R (1990) Description et cycle biologique d’ Hemolivia stellata n. g., n. sp., hémogrégarine de crapauds brésiliens. Annales de Parasitologie Humaine et Comparée 65:3–15. doi: 10.1051/parasite/1990651003

Siddall ME, Burreson EM (1994) The Development of a Hemogregarine of Lycodes raridens from Alaska in Its Definitive Leech Host. The Journal of Parasitology 80:569. doi: 10.2307/3283193 Introducción 37

Smith TG (1996) The genus Hepatozoon (Apicomplexa: Adeleina). J Parasitol 82:565–585

Telford SRJr (1988) A Contribution to the systematics of the reptilian malaria parasites, family Plasmodiidae (Apicomplexa Haemospororina). University of Florida, Gainesville

Telford SRJr (1996) Two new species of Haemocystidium Castellani & Willey (Apicomplexa: Plasmodiidae) from Pakistani lizards, and the support their meronts provide for the validity of the genus. Systematic Parasitology 34:197–214. doi: 10.1007/BF00009387

Telford SRJr (2009) Hemoparasites of the Reptilia: color atlas and text. Taylor & Francis, Boca Raton

Tomé B, Maia JPMC, Harris DJ (2012) Hepatozoon Infection Prevalence in Four Snake Genera: Influence of Diet, Prey Parasitemia Levels, or Parasite Type? Journal of Parasitology 98:913–917. doi: 10.1645/GE-3111.1

Valkiūnas G (2005) Avian malaria parasites and other haemosporidia. CRC Press, Boca Raton

Votýpka, J, Modrý, D., Oborník, M., et al (2017) Apicomplexa. In: Handbook of the Protists. Springer International Publishing., pp 1–58

Wenyon CM (1915) The pigmented parasites of cold-blooded animals, with some notes on a Plasmodium of the Trinidad iguana. Journal of Tropical Medicine and Hygiene 18:133–140

Wenyon CM (1926) Protozoology: a manual for medical men, veterinarians and zoologists. New York: W. Wood

IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

1. Capítulo 1. Plasmodium parasites in reptiles from the Colombia Orinoco- Amazon basin: a re-description of Plasmodium kentropyxi Lainson R, Landau I, Paperna I, 2001 and Plasmodium carmelinoi Lainson R, Franco CM, da Matta R, 2010 Parásitos de Plasmodium en reptiles de la cuenca Orinoco-Amazonas de Colombia: una nueva descripción de Plasmodium kentropyxi Lainson R, Landau I, Paperna I, 2001 y Plasmodium carmelinoi Lainson R, Franco CM, da Matta R, 2010.

Colombia es un país megadiverso con cerca de 600 especies de reptiles; sin embargo, hay pocos estudios sobre especies de hemoparásitos encontrados en este grupo taxonómico. Aquí, documentamos la presencia de Plasmodium spp. en cuatro especies de reptiles de la parte norte de la región Orinoco-Amazonas en Colombia. Los individuos analizados en este estudio fueron capturados en localidades entre 200 y 500 m de altitud, en el departamento de Guaviare. Cada muestra se analizó en busca de parásitos haemosporidios mediante el uso de una morfología y un protocolo de reacción en cadena de la polimerasa anidada (PCR) que se dirige al gen del citocromo b mitocondrial (cytb). Se encontraron cuatro morfotipos del género Plasmodium; dos de estas especies son re- descritas usando datos morfológicos y moleculares (cytb). Para los otros dos morfotipos, no fue posible asignar una especie descrita. Entre ellos, Plasmodium evaluado en una especie solo fue detectado por microscopía. Teniendo en cuenta la diversidad potencial de especies, es posible que los primers comúnmente utilizados no detecten todas las especies, lo que refuerza la importancia de usar microscopía en el análisis de hematozoos. No hubo correspondencia entre los rasgos morfológicos asociados con los subgéneros y las relaciones filogenéticas que encontramos en nuestros análisis. Además, encontramos una expansión en la distribución geográfica de estas dos especies, y un nuevo hospedero para P. kentropyxi, lo que demuestra que los estudios de herpetofauna tropical y sus parásitos merecen más atención. Capítulo 1 39

Palabras clave: Citocromo b; Hemoparásito; Herpetofauna; Neotropico; Plasmodium.

Este artículo abarca ampliamente los objetivos planteados en el proyecto de maestría, teniendo en cuenta que se estableció prevalencias, parasitemias y rango de hospedero, asi mismo, se realizó la determinación morfológica y molecular de dos especies de Plasmodium, identificados en reptiles de la región Orinoco-Amazonas en Colombia; además de asociarse estas dos caracterizaciones. En este artículo realicé en la lectura de los extendidos sanguíneos, la toma de fotografías y medidas de las formas parasitarias identificadas. Participé en la escritura del manuscrito inicial y las nuevas versiones solicitadas por los revisores. La versión presentada en este documento corresponde a la versión post print, por lo cual puede tener pequeñas diferencias con la versión publicada finalmente, que por derechos de la revista no puede ser cargada en esta tesis. La versión final puede ser consultada en Parasitology Research: https://link.springer.com/article/10.1007%2Fs00436-018-5815-9

40 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Plasmodium parasites in reptiles from the Colombia Orinoco-Amazon basin: a re-description of Plasmodium kentropyxi Lainson R, Landau I, Paperna I, 2001 and Plasmodium carmelinoi Lainson R, Franco CM, da Matta R, 2010

Nubia E. Matta 1 Leydy P. González 1,2 M. Andreína Pacheco 3 Ananías A. Escalante 3 Andrea M. Moreno 1,4 Angie D. González 1 Martha L. Calderón-Espinosa 5

1. Departamento de Biología, Facultad de Ciencias, Universidad Nacional de Colombia, Sede Bogotá, Carrera 30 No 45-03, Bogotá, Colombia. 2. Instituto de Biotecnología, Facultad de Ciencias, Universidad Nacional de Colombia, Sede Bogotá, Carrera 30 No 45-03, Bogotá, Colombia 3. Institute for Genomics and Evolutionary Medicine (IGEM), Temple University, Philadelphia, PA, USA 4. Universidad Colegio Mayor de Cundinamarca, Sede Bogotá, Calle 28 No 5B-02, Bogotá, Colombia 5. Instituto de Ciencias Naturales, Facultad de Ciencias, Universidad Nacional de Colombia, Sede Bogotá, Carrera 30 No 45-03, Bogotá, Colombia

ORCID

Nubia E. Matta: 0000-0003-1775-0804 Leydy P. González: 0000-0003-2494-0665 M. Andreína Pacheco: 0000-0002-5682-7299 Ananias A. Escalante: 0000-0002-1532-3430 Angie D. González: 0000-0002-1772-2152 Andrea M. Moreno: 0000-0002-4306-3613

Matta, N.E., González, L.P., Pacheco, M. et al. Parasitol Res (2018) 117: 1357. https://doi.org/10.1007/s00436-018-5815-9 Available in: https://link.springer.com/article/10.1007%2Fs00436-018-5815-9

Capítulo 1 41

1.1 Abstract Colombia is a megadiverse country with about 600 species of reptiles; however, there are few studies on species of hemoparasites found in this taxonomic group. Here, we document the presence of Plasmodium spp. in four species of reptiles from the northern part of the

Orinoco-Amazon region in Colombia. Individuals analyzed in this study were captured in localities between 200 and 500 m altitude, in the department of Guaviare. Each sample was screened for haemosporidian parasites by using morphology and a nested polymerase chain reaction (PCR) protocol that targets the mitochondrial cytochrome b (cytb) gene. Four morphotypes of the genus Plasmodium were found, two of these species are re-described using morphological and molecular data (cytb). For the other two morphotypes it was not possible to assign a described species. Among those Plasmodium screened one species was only detected by microscopy. Considering the potential species diversity, it is possible that commonly used primers may not detect all species, reinforcing the importance of using microscopy in haematozoa surveys. There was no correspondence between the morphological traits associated with the subgenera and the phylogenetic relationships that we found in our analyses. Additionally, we found an expansion in the geographical distribution of these two species, and a new host for P. kentropyxi, demonstrating that studies of tropical herpetofauna and their parasites deserve more attention.

Keywords: Cytochrome b, haemoparasite, herpetofauna, Neotropics, Plasmodium.

1.2 Introduction The Haemosporida comprise a group of cosmopolitan parasitic protozoa that infect mammals, birds, amphibians, and reptiles (Garnham 1966; Valkiūnas 2005). These vector- borne parasites undergo a complex life cycle with its sporogony (sexual stages followed by meiosis) in hematophagous dipterans, completing their asexual or schizogony (haploid 42 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA. stages) and dimorphic gametogony in their vertebrate hosts (Garnham 1966; Valkiūnas

2005; Telford S.R. Jr. 2009). Despite the extensive use of molecular methods, morphology remains the standard for the diagnosis and delimitation of species (Valkiūnas 2005; Telford

S.R. Jr. 2009). However, morphological intraspecific variability is still a problem. Parasite morphology varies during infection even in the same host species (Lainson et al. 2001).

Furthermore, there is increasing recognition of the importance of so-called cryptic species in parasitic organism (Nadler and De León 2011). Therefore, many researchers agree that the diagnosis from morphological features has serious limitations (Falk et al. 2011; Pacheco et al. 2013) giving rise to the determination of synonymous species, uncertain species or ambiguous designations (Perkins 2014; Outlaw and Ricklefs 2014). Nevertheless, since morphology remains the standard (Lainson et al. 2001; Telford S.R. Jr. 2009), there is a need for characterizing intraspecific variability.

Plasmodiidae is the most studied family within the Haemosporida since it includes the etiological agents of human malaria (Garnham 1966; Valkiūnas 2005). There are at least

100 recognized species that are parasites of reptiles; these species have been described based on morphological characters (Garnham 1966; Telford S.R. Jr. 1988). Garnham

(1966) proposed three subgenera within the family Plasmodiidae to classify the 23 known species in lizards up to that moment: Sauramoeba for parasites with large erythrocytic meronts, Carinamoeba for those with very small erythrocytic meronts, and the subgenus

Ophidiella for a Plasmodium species found in snakes. Later, Telford (1988) proposed seven subgenera for Plasmodium species infecting reptiles: Sauramoeba, Carinamoeba, Para

Plasmodium, Asiamoeba, Lacertamoeba, Garnia, and Ophidiella. This proposal is not free of controversy. Garnia is sometimes considered as a separate family, Garniidae, with three genera (Garnia, Fallisia, and Progarnia) due to the fact that these parasites do not produce hemozoin, and Fallisia parasites are only present in thrombocytes or leukocytes (Lainson Capítulo 1 43 et al. 1971, 1974; Valkiūnas 2005). In the case of Plasmodium in Neotropical lizards, there are at least 24 described species (Telford S.R. Jr. 1988) and they have been mainly studied in Brazil, México, and the Lesser Antilles. Only a few studies have been carried out in

Colombia. One of these studies reported the presence of Plasmodium floridense in Anolis concolor Cope, 1862 in the San Andrés and Providencia Islands and Plasmodium colombiense infecting A. auratus Daudin, 1802 in the valley of Rio Cauca (Ayala 1975;

Ayala and Spain 1976). Although Ayala (1978) refers to the presence of another six

Plasmodium species of reptiles in several colombian localities, there is no publication associated with those data.

The integration of morphological and molecular criteria to define species is emerging as a consensus in modern systematic parasitology (Nadler and De León 2011). Since its introduction in molecular phylogenetic studies, genes from the parasites’ mitochondria have strengthened the description of species (Escalante et al. 1998; Perkins and Schall 2002;

Ricklefs et al. 2004; Bensch et al. 2004). Indeed, the use of mitochondria has meant a significant increase in the knowledge of the genetic diversity, the discovery of new hostparasite-vector relationships, and a formal assessment of these parasites´ phylogenetic relationships (Escalante et al. 1998; Perkins and Schall 2002; Ricklefs et al. 2004; Sehgal et al. 2006; Martinsen et al. 2006; Pacheco et al. 2018). The use of molecular methods has allowed the characterization of cryptic and complex species such as the case of

Plasmodium azurophilum and Plasmodium leucocytica, initially described as a single species because of shared morphological characters (Telford S.R. Jr. 1975). However, a separation of their molecular lineages was possible, supported by the type of host cell present in each population (Perkins 2001; Telford S.R. Jr. 2009). 44 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

The present study aimed to document the presence of Plasmodium spp. in reptiles in the northern part of the Colombian Amazon region, re-describe the species found, and determine the association of molecular lineages with morphological Plasmodium species.

1.3 Materials and methods

1.3.1 Study area Reptiles were collected in the Guaviare department-Colombia in five localities between 200-

500 m altitude with an average annual temperature of 25-30 °C (Fig. I 1; Online Resource

1). This region represents a transition between the ecosystems of the eastern plains and the Amazonian forest and includes a variety of forests, natural savannas and rocky outcrops

(Escobar 2000).

Capítulo 1 45

1.3.2 Samples A total of 31 individuals belonging to 22 species (Table I 1) were captured and sacrificed with 2% lidocaine. Subsequently, a blood sample was taken from the portal vein and three slides were made, which were fixed with methanol for 5 min and stained with 10% Giemsa pH 7.2 for 45 min. Blood samples preserved in SET buffer (0.05 M Tris, 0.15 M NaCl, 0.5

M EDTA pH 8.0) were further stored for molecular analyses. Samples were kept at room temperature in the field and then cooled to -20 °C in the laboratory. The specimens were collected under a permit granted by the “Corporación para el Desarrollo Sostenible del

Norte y Oriente Amazónico-CDA” (CDA Resolution No. DSGV-112, October 25, 2012).

1.3.3 Microscopic analyses Blood slides were examined using a Leica DM750e microscope (Leica Microsystems,

Heerbrugg, Switzerland), at x400 for 10 min and then at x1000 for 20 min, and the slides with parasites were examined in their entirety for the capture of digital images, using a Leica

EC3 digital camera and processed with the LAS EZ (Leica Microsystems Suiza Limited

2012). Morphometric analysis was performed using ImageJ software (Schneider et al.

2012), and intensity of infection was estimated from erythrocyte counts with an increase of x1000, focusing on areas where blood cells formed a monolayer (No. of parasites/10,000 erythrocytes) (Staats and Schall 1996). The taxonomical determination of blood parasites was made comparing their morphologies with reptilian Plasmodiid species reported in

Lainson et al. (1975), Ayala and Spain (1976), Lainson et al. (2001), Telford S.R.Jr. (2009),

Lainson et al. (2010), and Lainson (2012). 46 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

1.3.4 Detection of haemosporidian parasites by polymerase chain reaction (PCR) Total genomic DNA was extracted from blood using QIAamp® DNA Blood Mini Kit (Qiagen

GmbH, Hilden, Germany). Each DNA sample was screened for haemosporidian parasites by using a nested polymerase chain reaction (PCR) that targets the mitochondrial cytochrome b (cyt b) gene in the parasite using the protocol described in Pacheco et al.

(2011). In addition to the cyt b gene sequences obtained in this study, partial mitochondrial genomes were also obtained for these parasites and published elsewhere (Pacheco et. al.

2018). However, for the purpose of this research, only the cyt b sequences obtained during the screening were analyzed, so we can compare with the data available from other studies.

All sequences included in our analyses are listed in Online Resource 2.

1.3.5 Phylogenetic analysis of the cyt b fragment Three independent nucleotide alignments were performed by using the MUSCLE algorithm

(Edgar 2004) implemented in SeaView4 (Gouy et al. 2010). The first alignment (500 bp. excluding gaps) was constructed with a total of 74 cyt b sequences of parasite species from lizards, birds, and mammals (ungulates, rodent, and primates) belonging to the genera

Leucocytozoon, Haemoproteus, Haemocystidium, and Plasmodium. For the second alignment (981 bp. excluding gaps) the short sequences of two lizard parasites P. zonuriae and P. intabazwe and the two sequences of ungulate Plasmodium were excluded. The third alignment (981 bp excluding gaps) was performed using only sequences of parasite species from lizards and birds. All sequences were translated into protein, and no amino acid insertion or deletion was found. Phylogenetic hypotheses were estimated by using

Bayesian Inference as implemented in MrBayes version 3.2.6 (Ronquist and Huelsenbeck

2003). The phylogenies were estimated using a General Time-Reversible model with gamma-distributed substitution rates and a proportion of invariable sites (GTR + Γ + I), Capítulo 1 47 which was the model that better fit the data with the lowest corrected Akaike Information

Criterion (AIC) scores as estimated by jModeltest 2.1.3 (Darriba et al. 2012). Two independent runs of 6x106 generations were conducted with four chains, sampling every five hundred generations in order to avoid auto-correlation. The chains were assumed to have converged once the average standard deviation of the posterior probability was below

0.01 and the value of the potential scale reduction factor (PSRF) was between 1.00 and

1.02 (Drummond et al. 2006) In total, 25% of the trees were discarded as a burn-in step.

1.4 Results Plasmodium spp. were encountered in four of the 31 (12.9%) samples examined and evaluated both by microscopic and molecular tools. Infections were reported in lizards belonging to the families Teiidae (n=2/3), Dactyloidae (n=1/5), and Tropiduridae (n=1/2)

(Table I 1; Online Resource 1). Despite Garnham (1966) and Telford (1988) reporting only the subgenus Ophidiella for Plasmodium infecting snakes, in the samples we analyzed here, we found no parasites similar to Ophidiella. Snakes were infected by Hepatozoon spp. that were not the subject of this study.

Table I 1. Reptiles sampled in this study and the prevalence of Plasmodium spp.

No. Individuals Family Species Parasite sampled/infected Lizards Cercosaura 1 argulus Gymnophtalmidae Leposoma 3 percarinatum Plasmodium Ameiva ameiva 1/1 (Carinamoeba) Teiidae carmelinoi Cnemidophorus 1 lemniscatus 48 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Plasmodium Cnemidophorus cf. 1/1 (Sauramoeba) gramivagus kentropyxi Plasmodium Anolis auratus 2/1 (Carinamoeba) sp. Anolis aff. auratus 1 Dactyloidae Anolis scypheus 1 Anolis 1 fuscoauratus Co-infection Plasmodium Tropiduridae Plica cf. plica 2/1 (Lacertamoeba) sp. and Schellakia sp. Lepidoblepharis Sphaerodactylidae 2 sp. Gonatodes riveroi 5 Scincidae Mabuya 1 altamazonica Snakes Ninia atrata 1 Atractus sp. 1 Helicops angulatus 1 Dipsadidae Imantodes 1 cenchoa Pseudoboa 1 coronata Mastigodryas Colubridae 1 boddaerti. Oxyrhopus 1 petolarius Boidae Corallus 1 hortulanus Epicatres chechria 1 Total 31 4

Morphological redescriptions

Plasmodium (Sauramoeba) kentropyxi (Lainson et al. 2001) (Fig I 2 a-f; Table I 2).

This parasite was first described in Kentropyx calcarata (Spix, 1825) from northern Brazil; here it was found infecting the lizard Cnemidophorus cf. gramivagus Mccrystal & Dixon,

1987 (Squamata: Teiidae). Gametocytes and meronts were found infecting both mature Capítulo 1 49 and immature erythrocytes. However, no trophozoites were observed. The host cell nucleus was displaced towards the periphery (Fig. I 2a-f), and also a notorious hypertrophy was observed (Table I 2, Fig. I 2c). The sex ratio (macro/microgametocytes) was 5:3. Double infections in the same erythrocyte were observed (Fig. I 2a). In the original description, such sex ratio was 4:1.

Meronts contain between 12-50 merozoites (Fig. I 2a-c). Mature meronts tend to occupy almost all the erythrocytes, displacing the nucleus of the host cell towards the poles and occasionally adhering to the nucleus. Immature meronts are in a lateral position and seem to grow and surround the nucleus; even in mature stages the nucleus has the appearance of being overlapped. Pigment granules are condensed into a single point clumping in a cytoplasmic vacuole, that has a mean diameter of 1.5 μm. All the above-mentioned features are shared with the original description (Lainson et al. 2001). It is important to mention that the Plasmodium kentropyxi reported here has a greater number of merozoites than in the original description (Table I 2).

Gametocytes were elongated as a band with rounded ends (Fig. I 2d-f) and the pigment granules were small and scattered and located in the cytoplasm. Sixty percent of the macrogametocytes (Fig. I 2e) showed at least one vacuole <1.1 μm. in diameter. However, they were not as conspicuous as described by Lainson et al. (2001), where the vacuole size was 2-3 μm. in diameter. Young macrogametocytes were located in the polar zone of the host cell and mature ones laterally and they did not attach to the nucleus. In immature stages of gametocytes the vacuole was not evident (Fig. I 2d). It is important to mention that macrogametocytes' nucleus was not as obvious as usual in another Plasmodium species (Fig. I 2e). This feature contrasts with the original description where the classical blue color of macrogametocytes was reported. Microgametocytes had a configuration similar to macrogametocytes with the pink coloration of the male (Fig. I 2f) having a diffuse 50 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA. nucleus and pigment granules distributed randomly in the cytoplasm. In the cytoplasm were observed until three small vacuoles (Fig. I 2f), similar to the original description of the vacuoles are located toward one edge of the parasite.

Taxonomic status

Host type: Kentropyx calcarata (Spix, 1825 -Teiidae) (Lainson et al. 2001) (Fig I 2g-l; Table I 2).

Parasite distribution: State of Pará in northern Brazil: Capanema (1° 12' S, 47° 11' W),

Belem (1° 27' S, 48° 29' W) and the Outeiro island (1° 18' S, 48° 28' W); new location

Guaviare, Colombia, (02º 30' 49.3'' S, 72º 42' 19.7'' W).

Additional host: Cnemidophorus cf. gramivagus Mccrystal & Dixon, 1987 (Teiidae)

Material: blood slides prepared for the present study were deposited in the collection of the

“Grupo de Estudio Relación Parásito-Hospedero (GERPH) at the Universidad Nacional de

Colombia under the collection code GERPH: Gu027.

DNA sequence: partial cyt b gene: MF177708; partial mtDNA: KY653753 (Pacheco et al. 2018).

Site of infection: mature erythrocytes, and immature erythrocytes at low frequency (5.4%).

Intensity of infection was 0.62

Additional material: None.

Remarks:

Plasmodium kentropyxi shares morphological characteristics with P. diploglossi (11-58 merozoites), P. guayannense (40-74 merozoites), P. achiotense (36-56 merozoites) and P. cnemidophori (42-127 merozoites), all of these are classified in the same subgenus

Sauramoeba. The main differential characteristics of Plasmodium kentropyxi are large meronts producing hypertrophy in the host cell and completely displacing the host cell Capítulo 1 51 nucleus. Moreover, the pigment granules are condensed in masses, and the parasite also has conspicuous vacuole(s) of variable size located at one end of the gametocytes. In comparison with P. diploglossi, that develops elongated meronts that encircle the host nucleus (Telford S.R. Jr. 2009), P. kentropyxi forms oval and roundish meronts that do not encircle the nucleus. In contrast, gametocytes of P. guayannense and P. achiotense are roundish, differing from the elongated band-like gametocytes formed by P. kentropyxi.

Plasmodium kentropyxi is also similar to P. cnemidophori, because their meronts cause hypertrophy and distortion of host cell and nucleus, as well as the displacement of the nucleus; moreover, some morphometric features of the taxa overlap. However, P. cnemidophori may infect proerythrocytes (Telford 1973), and produces on average more merozoites by meront, being 42-127 (Telford S.R Jr. 2009), while the average in P. kentropyxi is 12-50. Compared to P. pifanoi reported in Venezuela, it also has a conspicuous vacuole although there is a marked difference in the number of merozoites by meront. This latter species has only 7-16 merozoites, so this feature defines it as a species of the subgenus Paraplasmodium (Telford S.R. Jr. and S.R. Telford, III. 2003)

Plasmodium (Carinamoeba) carmelinoi (Lainson et al. 2010)

This taxon was found infecting the type host Ameiva ameiva Linnaeus, 1758; (Squamata:

Teiidae). Parasites were found infecting only mature erythrocytes located in a polar position in the host cell, and multiple infections were found in a single erythrocyte (data not shown).

Trophozoites were the most commonly found parasitic form (46% of the parasitic forms counted). They have cytoplasm that adopts a teardrop shape changing to oval or amoeboid when they grow. Dark browinish-black pigment granules tend to cluster over a conspicuous vacuole (Fig. I 2g). That vacuole increases in size as trophozoites develop. It is located at the center of the parasite and displaces the parasite cytoplasm towards the periphery. 52 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Meronts are mainly roundish and contain 6-10 small merozoites that do not adhere to the erythrocyte nucleus, thus, classifying this species in the subgenus Carinamoeba. Meronts have a large vacuole, that pushes the merozoites towards the periphery, giving a diadem appearance, (Fig. I 2h-k; Table I 2). Overall, the disposition and color of the pigment granules is similar to those described by Lainson et al. (2010); they are dark brownish-black pigment usually concentrating inside the vacuole (Fig I 2j). Only two out of 19 meronts observed, show a different organization (Fig. I 2k). In those cases, the pigment granules are more condensed, and the vacuole occupy less space in the parasitic cytoplasm and have a variable distribution of merozoites.

Gametocytes were observed at low frequency, limiting their characterization. The vacuole found in trophozoites and meronts was also observed in gametocytes with an oval shape and with a distal position in the cytoplasm of the parasite, also containing pigment granules

(Fig. I 2l) but smaller than in meronts. This characteristic was also observed by Lainson et al. (2010). Due to the low number of gametocytes observed it was not possible to determine the sex ratio, however, in the original description, this macro/microgametocyte ratio was

2.2:1.

Taxonomic status

Host type: Ameiva ameiva Linnaeus, 1758 (Lainson et al. 2010).

Parasite distribution: Pará in the Brazilian Amazon region, the state of Pará (01° 12' N, 47°

11' W). The new locality is the department of Guaviare in the northern Colombian Amazon (02º 27' 56.9'' N,

72º 42' 29.1'' W).

Additional host: None known Capítulo 1 53

Material: blood slides prepared for the present study were deposited in the biological collection GERPH of the Universidad Nacional de Colombia under the code GERPH:

Gu040.

DNA sequence: partial cyt b gene: MF177709; partial mtDNA: KY653755 (Pacheco et al.

2018).

Site of infection: mature erythrocytes

Intensity of infection was 0.15%

Remarks:

Differential diagnosis of this parasite should be made with P. attenuatum (4-8 merozoites),

P. diminutivum (4-6 merozoites), P. vacuolatum (8-20-merozoites), and P. attenuatum that are found infecting the same host in Guyana and Venezuela. These species differ from P. carmelinoi due to the presence of elongated gametocytes and rosette or fan-shaped meronts. In comparison with P. diminutivum (reported also in the same host in Panama) both parasite species have roundish gametocytes. However, P. diminutivum has meronts without the characteristic vacuole. Plasmodium carmelinoi and P. vacuolatum (Lainson et al. 1975), are very similar in size and both possess a prominent vacuole; they can be distinguished because P. carmelinoi has meronts with crown-shaped merozoites arranged around the vacuole. Meronts and gametocytes of P. carmelinoi are mainly round, whilst in

P. vacuolatum, they have the form of a bean. Finally, P. carmelinoi has an amoeboid form in the trophozoites stages (the main form found here), while P. vacuolatum does not.

Plasmodium sp.

This morphotype was found in the lizard Anolis auratus Daudin, 1802. 54 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Trophozoites were observed in mature erythrocytes with a pigment granule with size >2

μm, they are rounded and located variably in the cytoplasm (Fig. I 2m).

Meronts were oval or irregular in shape and had 2-12 merozoites irregularly arranged or in a fan shape. Pigment granules were clumped in a golden mass that was located in the peripherical zone (Fig. I 2n-o). Eighty-one percent of the meronts were immature

Gametocytes are oval in form and are polar in the erythrocyte; their size always is smaller than the host nucleus; the pigment granule arrangement is variable and dispersed in the cytoplasm or located in masses (Fig. I 2p). Vacuoles when present have a variable size.

Taxonomic status

Host: Anolis auratus Daudin, 1802

Locality: Guaviare, Colombia, (02º 30'49.3'' N, 72º 42' 19.7'' W).

Material: blood smears were deposited in the collection GERPH of the Universidad Nacional de Colombia, code GERPH: Gu030.

DNA sequence: It was not possible to amplify cyt b gene.

Site of infection: All parasite forms observed were infecting mature erythrocytes.

Intensity of infection was 0.1%.

Plasmodium (Lacertamaoeba) sp.

This Plasmodium infects mature erythrocytes of individuals from Plica cf. plica Linnaeus,

1758, appearing as a coinfection with a morphotype associated with Schellackia cf. sp.,

(result is not shown).

Trophozoites were rounded and had one pigment granule.

Meronts were present with 7-15 merozoites, located in a polar position, rounded- or oval- shaped with merozoites arranged irregularly or less frequently in a fan (Fig. I 2q-r). In young meronts there was a small vacuole (Fig. I 2q). Capítulo 1 55

Gametocytes were observed at low frequency, with roundish or oval forms that adhered to the nucleus of the erythrocyte (Fig. I 2s-t). In both macro and microgametocytes a variable distribution of the pigment granules was observed with some clusters. They were preferentially visible in polar position, and one vacuole with a diameter <4 μm was observed in some gametocytes (Fig. I 2t).

Taxonomic status

Host: Plica cf. plica Linnaeus, 1758

Locality: Guaviare, Colombia. (02º 27' 56.9'' N, 72º 42' 29.1'' W), 200-500 m a.s.l

Material: blood smears were deposited in the biological collection GERPH of the

Universidad Nacional de Colombia, under code GERPH: Gu010.

DNA sequence: partial cyt b gene: MF177707; partial mtDNA: KY653796 (Pacheco et al.

2018).

Site of infection: Mature erythrocytes;

Intensity of infection was 0.12% 56 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Capítulo 1 57

Plasmodium Plasmodium Plasmodium Plasmodium Characteristic (Sauramoeba) (Carinamoeba) (Carinamoeba) (Lacertamoeba) sp. kentropyxi carmelinoi sp. Non infected n = 37 n = 10 n = 30 n = 10 erythrocytes 13.5 ± 1.2 16,56 ± 0.84 15 ± 0.6 15,98 ± 1,81 Large (13,54 – 19,42) (15,34 – 17,94) (11.4 - 16.8) (13,7 – 19,4) 8.4 ± 0.87 10,06 ± 0,71 8.7 ± 0.5 11,29 ± 1,34 Wide (5,73 – 9,95) (8,83 – 11,19) (6.9- 10.9) (8,77 – 12,69) 115.2 ± 15.3 132,52 ± 9,69 103.6 ± 9.3 149,99 ± 17,86 Area (80,53 – 139,33) (118,53 – 148,06) (83.9- 131.2) (126,84 – 185,18) Nucleus Uninfected erythrocytes 5.8 ± 0.6 7,02 ± 0,44 6.4 ± 0.5 6,72 ± 0,30 Large (5,06 – 8,87) (6,44 – 7,75) (5.2- 6.9) (6,20 – 7,04) 4.2 ± 0.5 4,68 ± 0,36 3.7 ± 0.3 4,49 ± 0,32 Wide (3,17 – 5,38) (4,17 – 5,22) (3.1-6.5) (4,14 – 5,028) 21.1 ± 3.2 26,94 ± 2,68 19.6 ± 2.3 24,56 ± 1,20 Area (15,13 – 27,00) (23,32 – 30,42) (15.3-21.3) (22,08 – 26,07) Trophozoite / n = 2 n = 19 n = 3 3,65 ± 0,69 9.4 ± 2.5 3,76 ± 0,64 Area / (3,17 – 4,14) (6.2- 15.7) (2,75 – 5,01) 6.8 ± 1.8 Vacuole area / / / ( 4.3-11.8) Meronts n = 21 n= 4 n = 18 n = 7 9.3 ± 1.9 4,22 ± 0,84 4.9 ± 0.5 3,96 ± 0,78 Large (13.5 – 6.1) (44,22 – 6,13) (4.0-5.7) (2,89 – 5,017) 7 ± 2.1 6,24 ± 1,85 4.1 ± 0.7 3,96 ± 1,85 Wide (10.2 – 2.6) (3,7 – 7,98) (3.1-5.7) (1,64 – 7,63) 55 ± 22.1 28,44 ± 7,46 14.86 ± 2.07 13,66 ± 6,27 Area (87.5 - 14.9) (19,69 – 37,83) (11.1 -19.8) (7,48 – 26,43) 28.6 ± 18.5 No. of merozoits 6 - 12 5-6 2 – 9 (53 - 2) Macrogametocyte n = 10 n = 1 n = 3 n = 2 14.1 ± 3.9 4.365 ± 0.24 4.99 ± 0.22 Large 6.48 (18.2 – 6.7) (4.19 – 4.54) (4.83 – 5.14) 4.2 ± 1.1 4.06 ± 0.70 3.83 ± 0.22 Wide 4.36 (6.2 – 2.7) (3.56 – 4.56) (3.67 – 3.83) 55.9 ± 22.5 13.60 ± 0.43 17.25 ± 2.18 Area 24.42 (83.7 – 27.9) (13.3 – 13.91) (15.70 – 18.79) N. máximo de gránulos 7.4 ± 7.03 Accumulated Accumulated 7 – 6 de Pigment (22 – 0) Microgametocytes n = 7 n = 1 n = 1 n = 2 13.2 2.9 7.14 ± 0.067 Large 9.5 4.88 (16.5 – 8.6) (7.096 – 7.096) 5 ± 2.2 4.22 ± 0.27 Wide 4.73 3.79 (8.7 – 2.9) (4.030 – 4.42) 57.8 ± 15.6 24.61 ± 0.57 Area 9.5 14.97 (88.3 – 45) (24.21 – 25.02) Nº. Maximum of 10.57 ± 3.30 4 7 10 pigment granules (6 - 15) Accumulated

58 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

1.4.1 Molecular phylogenies The topologies of the three estimated phylogenies using the partial cyt b sequences (500 and 981 bp. excluding gaps) were consistent for many clades; however, some discrepancies can be observed (Fig. I 3 and Fig. I 4, Online Resource 3). Some phylogenetic relationships changed depending on the sequence length, but many of those clades were not supported due to their low posterior probabilities. Overall, the genera

Haemocystidium and Plasmodium were monophyletic groups sharing a common ancestor with Haemoproteus (Parahaemoproteus) species. However, Plasmodium species of lizards were not monophyletic. Indeed, depending on the tree, rodent and primate Plasmodium species shared a common ancestor with P. agamae and P. giganteum (Fig. I 3) or P. mexicanum and P. chiricahua (Online Resource 3). In addition, P. zonuriae and P. intabazwe also group with some bird Plasmodium parasites (Fig. I 3). Although these results may suggest host switches between lizards-birds and lizards-mammals, the low posterior probabilities in some of those clades indicate that a more robust phylogeny (e.g. multiple nuclear loci or complete mtDNA) is needed before reaching conclusions; however, such comprenhensive data is not yet available for reptile parasites.

Genetic distances on the cytb between all lizards Plasmodium species are shown in Online

Resource 4. The genetic distances obtained in this study indicated that all of the divergences are consistent with different species. In the case of the three lineages reported here, P. kentropyxi and P. carmelinoi always grouped together and shared a common ancestor with P. azurophilum and P. fairchildi (Fig. I 3, Fig. I 4, Online Resource 3, and

Online Resource 4), parasites that infect several species of lizard (genus Anolis) from the

Caribbean islands (Telford S.R. Jr. 2009). The lineage Plasmodium (Lacertamoeba) sp. found in this study and P. (Lacertamoeba) floridense belong to a different clade, sharing a common ancestor with P. (Lacertamoeba) hispaniolae (posterior probability = 1.0; Fig. I 3, Capítulo 1 59

4, Online Resource 3). Interestingly, all three parasites are from the Neotropics and belong to the subgenus Lacertamoeba. However, they were not monophyletic with other African P.

(Lacertamoeba) species (e.g., P. agamae). Indeed, given the phylogenies estimated in this study, none of the subgenera were monophyletic groups.

Figure I 3 A Bayesian phylogenetic hypothesis of lizard haemosporidian parasites based on partial sequences of cyt b gene (74 sequences and 500 bp excluding gaps). The values above branches are posterior probabilities (see Material and Methods). Leucocytozoon genus was used as outgroup. 60 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

1.5 Discussion Most Plasmodium species described thus far are found infecting birds and reptiles

(Valkiūnas 2005; Telford S.R. Jr. 2009). However, we are just starting to assess their diversity. Classical studies focused solely on morphological descriptions with limited information on the variation of the characters used or their capacity to distinguish species in a broad range of hosts (Falk et al. 2011; Pacheco et al. 2013). In addition, there are only a few morphotypes of those species associated with suitable molecular information that facilitates their identification. In the context of malarial parasites, the introduction of mitochondrial genes in phylogenetic investigations in the late 90’s (Escalante et al. 1998) lead to extensive data on the cyt b that has the potential to be used for barcoding (Outlaw and Ricklefs 2014). This proposal gained support in a recent study with well-characterized species where it was concluded that cyt b correctly separated the species in a sampling with more than 100 species of haemosporidia parasites (Pacheco et al. 2018). However, the taxonomic sampling linking parasite species from birds and reptiles with molecular lineages is still a work in progress. Paucity of data is particularly staggering in saurian malarias (see Pacheco et al 2018 where only 6 complete mitochondrial genomes have been published). Only twenty out of the at least 100 morphological species of Plasmodium infecting reptiles have a cyt b molecular lineage associated (including this study). Here we report three Plasmodium species, belonging to three of the seven subgenera reported in reptiles. Based on their morphology, we identify those species as P. (Sauramoeba) kentropyxi, P. (Carinamoeba) camelinoi, and P. (Lacertamoeba) sp. Given the limited taxonomic sampling, we observed no noticeable differences with the phylogenetic relationships found by Perkins and Austin (2009) and Falk et al. (2011) here. In contrast,

Pacheco et al 2018 found Plasmodium of reptiles as part of a monophyletic clade. Such difference could be explained, at least in part, by the scarce number mtDNA genomes 62 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA. available from these reptile parasites when compared to species with cyt b data.

Considering that only twenty species out of a hundred described have molecular data, it is evident that we need more studies in order to understand the evolution of these parasites.

In particular, it is pressing to link morphological and molecular data as is being done with avian parasites.

It is worth mentioning that our phylogenetic analyses, using partial cyt b seuqences, do not support the taxonomic conclusions of the subgenera classification. e.g., although P.

(Sauramoeba) kentropyxi and P. (Carinamoeba) carmelinoi appear as a sister taxa, they belong to different subgenera (Fig I 3, 4). Similar results have been found in Plasmodium subgenera of birds (Martinsen et al. 2007; Pacheco et al. 2018).

Although the geographic distribution of the host species for Plasmodium kentropyxi and P. carmelinoi overlaps in the northern part of South America, this report added new hosts and localities for these parasite species. These species shared a common ancestor with P. azurophilum and P. fairchildi, both species from the Caribbean Island (Ayala and Hertz

1981; Telford S.R. Jr. 2009). However, the taxonomic sampling is still too limited for further speculation on other patterns. Nevertheless, the four species in this clade have different morphological features, and overall, there is no correspondence between morphological traits and the actual phylogenetic relationships found in our analyses (Figs.I 3, 4). Although the number of parasites analyzed in our samples was fewer than those evaluated in original descriptions, we found parasites with a greater number of merozoites (Table I 2). Lainson et al. 2001 found a minimum effect of meronts on the host cell; this is different from our samples where the erythrocytes infected were considerable larger thanuninfected ones

(Table I 2), probably because of their greater number of merozoites. Capítulo 1 63

The lineage Plasmodium (Lacertamoeba) sp. found here infecting Plica cf. plica, has morphological features shared with P. floridense (4-32 merozoites), P. hispananiolae (4-8 merozoites), P. vacuolatum (8-20 merozoites), and P. colombiense (4-14 merozoites). In the phylogenetic analyses, Plasmodium (L.) sp. was grouped with P. (L.) floridense, a species reported in North America, Central America, and the Caribbean Islands, as well as in Colombia in northern South America (Telford Jr S.R 2009). Indeed, P. floridense and their different subspecies are very similar to Plasmodium (L.) sp., mainly because of their rosette-shaped meronts and the ovoid to elongated forms of their gametocytes. However, the specimen found in this study showed smaller size than those reported for P. floridense.

In addition, Plasmodium (L.) sp. and P. floridense shared a common ancestor with

Plasmodium hispaniolae (Fig. I 3, 4), which is also distributed in the Caribbean.

Morphological characteristics of this parasite are quite similar to P. floridense; however, one difference between these two species is that P. hispaniolae has mature gametocytes often with irregular and ragged cell margins. But the great majority of morphometric measurements overlap with P. floridense (Falk et. al. 2011). Plasmodium (L.) sp. is also morphologically very similar to P. hispaniolae, mainly because of the oval shape of gametocytes with vacuole, shape, and organization of their meronts. However, Plasmodium

(L.) sp. meronts and gametocytes are smaller than that reported for P. (L.) hispaniolae and this could be an effect of the host, as has been reported in the Plasmodium of birds (Mantilla et al. 2013).

Plasmodium (L.) sp. also has morphological characteristics similar to P. (L.) vacuolatum found in lizards from South America, but the vacuoles observed in our samples are not as conspicuous as the original description of P. vacuolatum (Lainson et al. 1975).

Unfortunately, there is no cyt b sequence available for P. vacuolatum that allows us to confirm a close phylogenetic relationship with Plasmodium (L.) sp. Finally, another species 64 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA. morphologically similar to Plasmodium (L.) sp. that has been reported for Colombia is P.

(L.) colombiense. Both species share the shape of gametocytes and the presence of vacuoles (less conspicuous in P. colombiense), but the distribution of pigment granules and size of gametocytes differs between them (Ayala and Spain 1976).

In this study, the host Plica cf. plica infected with Plasmodium (L.) sp. also was infected with a morphotype associated with the hemogregarina identified as Schellackia cf. sp.

(Bonorris and Ball 1955) (data not shown). Lizards of the genus Plica (Tropiduridae) are characterized by having a scansorial habit usuallyfound in forests, on vertical rocks and on trunks, and with a wide distribution towards the east of the Andes (Murphy and Jowers

2013). These lizards have frequently been reported infested by a large variety of mites

(Gomides et al. 2015) that could be vectors (Bonorris and Ball 1955), so these characteristics may explain the mixed infection found here.

The parasite found in Anolis auratus has a very particular arrangement of meronts, but due to the low intensity of infection and the lack of a cyt b sequence, we only attempt to identify it as Plasmodium sp. Overall this parasite has small meronts and the size of the gametocytes are similar to those described for parasites of the subgenus (Carinamoeba).

However, several stages have measurements that overlap with sizes described for

Lacertamoeba parasites (Table I 2) and for that reason we were not able to assign a definite subgenus. Interesting Anolis is a common genus infected by saurian malaria, e.g., the six malaria species described in the Caribbean Islands infect anoles (Anolis spp.) (Falk et al.

2011). Additionally, anoles also have been found infected by P. balli, P. marginatum and P. colombiense among others (Ayala and Hertz 1981; Perkins 2000). Ameiva ameiva is also a common host of a great variety of hemoparasites, including other Plasmodium species,

Hemolivia and Garnia (Lainson et al. 2003, 2010). It is a conspicuous lizard of the Teiidae Capítulo 1 65 family that has one of the widest geographic distribution among New World lizards, occurring from Panamá, Caribbean Island to Brazil (Avila-Pires 1995), this species occurs in a great array of ecosystems, but apparently it has an affinity for open microhabitats (Vitt and Colli 1994).

In relation to the impact of infection by Plasmodium spp. in reptiles, there are contrasting results; some authors consider them to be non-pathogenic (Brown et al. 2006; Stacy et al.

2011; Xuereb et al. 2012). However, negative effects have also been reported such as reduced fertility, limited feeding, reduced body mass, effects on liver and kidneys, anemia and variation in leukocyte parameters, possibly increasing susceptibility to secondary infections and predation, among other effects (Schall 2002; Vardo-Zalik and Schall 2008;

Motz et al. 2014). We did not follow the reptiles for a long period, so we have no data on this topic.

There are important gaps in our knowledge of blood parasites that infect wild species.

Specifically, in reptiles our understanding is fragmentary, where the great majority of species only have morphological characteristics; and almost certainly the species diversity in reptiles is underestimated, as it has been reported for avian hemoparasites. Although

PCR amplification is more sensible than microscopic analyses, those essays have not been properly standardized in many parasite species. Indeed, we were able to detect a parasite identified as Plasmodium sp. (Fig. I 2n-t) only by microscopy, despite also testing a different set of primers (e.g., Hellgren et al. 2004; Waldeström et al. 2004) and variations in the protocols with a gradient of Tm (data not shown). Our results provide support to Garnham´s

(1966) statements regarding the limits of morphological characteristics in saurian parasites: their number of merozoites is unstable, and the size of the parasite may depend on the host; for those reasons, it is not easily assigned a species or even a subgenera group.

Additional information concerning the biology of the parasites, their hosts, and their 66 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA. ecological relationships is necessary for an adequate approximation in the hemoparasite species determination in reptiles.

Knowing adequate baseline information about which parasites and which hosts’ species are in an ecosystem allow the characterization of trophic relationships and knowledge of possible reservoirs of potential pathogens, elements that make a difference in the design of appropriate conservation programs. Lizards like Ameiva ameiva or Anolis spp. would expand to new habitats and with them their parasites. As more forested habitats will be converted into pasturelands contact of naïve hosts with new parasites could cause a loss in biodiversity, as has occurred in avian malaria (van Riper III et al. 1986; Alley et al. 2010).

Conflict of interest: The authors declare they have no conflict of interest.

1.6 Acknowledgements The authors wish to thank the local community organization of the Playa Guio Natural

Reserve, San José del Guaviare for providing facilities for field logistics. We thank the students of the course "Animal Systematics" for their help in fieldwork. Professor Thomas

Defler kindly corrected the English of this manuscript. This study was partially supported by the “Dirección de Investigación” of the Universidad Nacional de Colombia, Bogota (Project

22940); and by the “Departamento Administrativo de Ciencias, Tecnologia e Innovacion”

COLCIENCIAS (Project code 1101-776-57872). The funding source had no role in study design, data collection, and analysis, or preparation of the manuscript. We thank the DNA laboratory at the School of Life Sciences (Arizona State University) for their technical support. Capítulo 1 67

1.7 Reference Alley M, Hale K, Cash W, et al. (2010) Concurrent avian malaria and avipox virus infection in translocated South Island saddlebacks (Philesturnus carunculatus carunculatus). N Z Vet J 58:218–223. doi: 10.1080/00480169.2010.68868

Avila-Pires TC. (1995) Lizard of Brazilian Amazonia (Reptilia: Squamata) Ayala SC (1975) Malaria and hemogregarines from lizards of the Western Caribbean Islands of San Andrés and Providencia. Rev Inst Med Trop São Paulo 17:218–224

Ayala SC (1978) Checklist, Host Index, and Annotated Bibliography of Plasmodium from Reptiles. J Protozool 25:87–100. doi: 10.1111/j.1550-7408.1978.tb03874.x

Ayala SC, Hertz PE (1981) Malaria infection in Anolis lizards on Martinique, Lesser Antilles. Rev Inst Med Trop Sao Paulo 23:12–17

Ayala SC, Spain JL (1976) A population of Plasmodium colombiense sp. n. in the iguanid lizard, Anolis Auratus. J Parasitol 62:177–189

Bensch S, Pérez-Tris J, Waldenström J, Hellgren O (2004) Linkage between nuclear and mitochondrial DNA sequences in avian malaria parasites: multiple cases of cryptic speciation? Evol Int J Org Evol 58:1617–1621

Bonorris JS, Ball GH (1955) Schellackia occidentalis n.sp., a Blood-inhabiting Coccidian Found in Lizards in Southern California. J Protozool 2:31–34. doi: 10.1111/j.1550- 7408.1955.tb02393.x

Brown GP, Shilton CM, Shine R (2006) Do parasites matter? Assessing the fitness consequences of haemogregarine infection in snakes. Can J Zool 84:668–676. doi: 10.1139/z06-044

Darriba D, Taboada GL, Doallo R, Posada D (2012) jModelTest 2: more models, new heuristics and parallel computing. Nat Methods 9:772–772. doi: 10.1038/nmeth.2109

Drummond AJ, Ho SYW, Phillips MJ, Rambaut A (2006) Relaxed Phylogenetics and Dating with Confidence. PLoS Biol 4:e88 . doi: 10.1371/journal.pbio.0040088

Edgar RC (2004) MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Res 32:1792–1797 doi: 10.1093/nar/gkh340

Escalante AA, Freeland DE, Collins WE, Lal AA (1998) The evolution of primate malaria parasites based on the gene encoding cytochrome b from the linear mitochondrial genome. Proc Natl Acad Sci U S A 95:8124–8129

Escobar S. F (2000) Diversidad de coleopteros coprofagos (Scarabaeidae: Scarabaeinae) en un mosaico de habitats en la reserva natural Nukak, Guaviare, Colombia. Acta Zool Mex 103–121 68 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Falk BG, Luke Mahler D, Perkins S (2011) Tree-based delimitation of morphologically ambiguous taxa: A study of the lizard malaria parasites on the Caribbean island of Hispaniola. Int J Parasitol 41:967–980. doi: 10.1016/j.ijpara.2011.05.004

Garnham PCC (1966) Malaria Parasites and other Haemosporidia. Blackwell Scientific Publications, Oxford, U. K

Gomides SC, Maturano R, Daemon E, et al. (2015) New reports of Acari ectoparasites on lizards of the genus Plica (Squamata: Tropiduridae) and a list of parasites known from this genus. Deutsche Gesellschaft Herpetologie Terrarienkunde ev c/o Jorn Kohler, Hessisches Landesmuseum Darmstadt, Friedensplatz 1, Darmstadt, 64283, Germany

Gouy M, Guindon S, Gascuel O (2010) SeaView Version 4: A Multiplatform Graphical User Interface for Sequence Alignment and Phylogenetic Tree Building. Mol Biol Evol 27:221–224 . doi: 10.1093/molbev/msp259

Hellgren O, Waldeström J, Bensch S (2004) A new PCR assay for simultaneous studies of Leucocytozoon, Plasmodium, and Haemoproteus from avian blood. J Parasitol 90:797–802

Lainson R (2012) Atlas de parasitas protozoários da fauna da Amazônia Brasileira: Haemosporida de répteis. Instituto Evandro Chagas, Ananindeua

Lainson R, Franco CM, da Matta R (2010) Plasmodium carmelinoi n. sp. (Haemosporida: Plasmodiidae) of the lizard Ameiva ameiva (Squamata: Teiidae) in Amazonian Brazil. Parasite 17:129–132. doi: 10.1051/parasite/2010172129

Lainson R, Landau I, Paperna I (2001) Plasmodium kentropyxi n.sp. (Apicomplexa: Haemosporina: Plasmodiidae) and a Plasmodium tropiduri-like parasite in the lizard Kentropyx calcarata (Lacertilia: Teiidae) in north Brazil. Parasite Paris Fr 8:107–113

Lainson R, Landau I, Shaw JJ (1974) Observations on non-pigmented haemosporidia of Brazilian lizards, including a new species of Saurocytozoon in Mabuya mabouya (Scincidae). Parasitology 69:215–223

Lainson R, Landau I, Shaw JJ (1971) On a new family of non-pigmented parasites in the blood of reptiles: Garniidae fam. nov., (Coccidiida: Haemosporidiidea). Some species of the new genus Garnia. Int J Parasitol 1:241–250. doi: 10.1016/0020- 7519(71)90027-0

Lainson R, Shaw JJ, Landau I (1975) Some blood parasites of the Brazilian lizards Plica umbra and Uranoscodon superciliosa (Iguanidae). Parasitology 70:119–141. doi: 10.1017/S0031182000048927 Capítulo 1 69

Lainson R, Souza MC de, Franco CM (2003) Haematozoan parasites of the lizard Ameiva ameiva (Teiidae) from Amazonian Brazil: a preliminary note. Mem Inst Oswaldo Cruz 98:1067–1070. doi: 10.1590/S0074-02762003000800016

Leica Microsystems Suiza Limited (2012) Leica Microsystems Suiza Limited. In: Leica Microsyst. http://www.leica-microsystems.com/products/microscope- software/software-for-materials-sciences/details/product/leica-application-suite/. Accessed 7 Jul 2015

Nadler SA, De León GP (2011) Integrating molecular and morphological approaches for characterizing parasite cryptic species: implications for parasitology. Parasitology 138:1688-1709. doi: 10.1017/S003118201000168X.

Mantilla JS, González AD, Valkiūnas G, et al. (2013) Description and molecular characterization of Plasmodium (Novyella) unalis sp. nov. from the Great Thrush (Turdus fuscater) in highland of Colombia. Parasitol Res 112:4193–4204

Martinsen ES, Paperna I, Schall JJ (2006) Morphological versus molecular identification of avian Haemosporidia: an exploration of three species concepts. Parasitology 133:279 . doi: 10.1017/S0031182006000424

Martinsen ES, Waite JL, Schall JJ (2007) Morphologically defined subgenera of Plasmodium from avian hosts: test of monophyly by phylogenetic analysis of two mitochondrial genes. Parasitology 134:483. doi: 10.1017/S0031182006001922

Motz VL, Lewis WD, Vardo-Zalik AM (2014) Leukocyte Profiles for Western Fence Lizards, Sceloporus occidentalis, Naturally Infected by the Malaria Parasite Plasmodium mexicanum. J Parasitol 100:592–597. doi: 10.1645/13-371.1

Murphy J, Jowers M (2013) Treerunners, cryptic lizards of the Plica plica group (Squamata, Sauria, Tropiduridae) of northern South America. ZooKeys 355:49–77. doi: 10.3897/zookeys.355.5868

Outlaw DC, Ricklefs RE (2014) Species limits in avian malaria parasites (Haemosporida): how to move forward in the molecular era. Parasitology 141:1223–1232. doi: 10.1017/S0031182014000560

Pacheco MA, Cranfield M, Cameron K, et al. (2013) Malarial parasite diversity in chimpanzees: the value of comparative approaches to ascertain the evolution of Plasmodium falciparum antigens. Malaria J. 12:328. doi: 10.1186/1475-2875-12-328.

Pacheco MA, Escalante AA, Garner MM, et al (2011) Haemosporidian infection in captive masked bobwhite quail (Colinus virginianus ridgwayi), an endangered subspecies of the northern bobwhite quail. Vet Parasitol 182:113–120. doi: 10.1016/j.vetpar.2011.06.006 70 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Pacheco MA, Matta NE, Valkiunas G, et al. (2018) Mode and rate of evolution of haemosporidian mitochondrial genomes: timing the radiation of avian parasites. Mol Biol Evol. 35(2):383-403. doi: 10.1093/molbev/msx285

Perkins SL (2001) Phylogeography of Caribbean lizard malaria: tracing the history of vector- borne parasites. J Evol Biol 14:34–45

Perkins SL (2000) Species concepts and malaria parasites: detecting a cryptic species of Plasmodium. Proc R Soc B Biol Sci 267:2345–2350

Perkins SL (2014) Malaria’s Many Mates: Past, Present, and Future of the Systematics of the Order Haemosporida. J Parasitol 100:11–25. doi: 10.1645/13-362.1

Perkins SL, Austin CC (2009) Four new species of Plasmodium from New Guinea lizards: integrating morphology and molecules. J Parasitol 95:424–433

Perkins SL, Schall J (2002) A molecular phylogeny of malarial parasites recovered from cytochrome b gene sequences. J Parasitol 88:972–978

Ricklefs RE, Fallon SM, Bermingham E (2004) Evolutionary relationships, cospeciation, and host switching in avian malaria parasites. Syst Biol 53:111–119

Ronquist F, Huelsenbeck JP (2003) MrBayes 3: Bayesian phylogenetic inference under mixed models. Bioinforma Oxf Engl 19:1572–1574

Schall JJ (2002) Parasite Virulence. Behav. Ecol. Parasites 283–313

Schneider CA, Rasband WS, Eliceiri KW (2012) NIH Image to ImageJ: 25 years of image analysis. Nat Methods 9:671–675. doi: 10.1038/nmeth.2089

Sehgal RNM, Hull AC, Anderson NL, et al. (2006) Evidence for cryptic speciation of Leucocytozoon spp. (Haemosporida, Leucocytozoidae) in diurnal raptors. J Parasitol 92:375–379

Staats CM, Schall JJ (1996) Malarial Parasites (Plasmodium) of Anolis Lizards: Biogeography in the Lesser Antilles. Biotropica 28:388–393. doi: 10.2307/2389202

Stacy NI, Alleman AR, Sayler KA (2011) Diagnostic Hematology of Reptiles. Clin Lab Med 31:87–108 . doi: 10.1016/j.cll.2010.10.006

Telford SR Jr. (1973) Malaria Parasites of the “Borriguerro’’Lizard, Ameiva ameiva (Sauria: Teiidae) in Panama. J Protozool 20:203–207. doi: 10.1111/j.1550- 7408.1973.tb00863.x

Telford S.R. Jr. (1975) Saurian malaria in the Caribbean: Plasmodium azurophilum sp. nov., a malarial parasite with schizogony and gametogony in both red and white blood cells. Int J Parasitol 5:383–394. doi: 10.1016/0020-7519(75)90003-X Capítulo 1 71

Telford S.R. Jr. (1988) A Contribution to the systematics of the reptilian malaria parasites, family Plasmodiidae (Apicomplexa Haemospororina). University of Florida, Gainesville

Telford S.R. Jr. (2009) Hemoparasites of the Reptilia: color atlas and text. Taylor & Francis, Boca Raton

Telford S.R Jr., S. R. Telford, III. (2003) Rediscovery and redescription of Plasmodium pifanoi and description of two additional Plasmodium parasites of Venezuelan lizards. J Parasitol 89:362–368. doi: 10.1645/0022-3395(2003)089[0362: RAROPP]2.0.CO;2

Valkiūnas G (2005) Avian malaria parasites and other haemosporidia. CRC press van Riper III C, van Riper SG, Goff ML, Laird M (1986) The Epizootiology and Ecological Significance of Malaria in Hawaiian Land Birds. Ecol Monogr 56:327. doi: 10.2307/1942550

Vardo-Zalik AM, Schall JJ (2008) Clonal diversity within infections and the virulence of a malaria parasite, Plasmodium mexicanum. Parasitology 135:1363–1372. doi: 10.1017/S0031182008004964

Vitt LJ, Colli GR (1994) Geographical ecology of a Neotropical lizard: Ameiva ameiva (Teiidae) in Brazil. Can J Zool 72:1986–2008. doi: 10.1139/z94-271

Waldenström J, Bensch S, Hasselquist D, Ostman O (2004) A new nested polymerase chain reaction method very efficient in detecting Plasmodium and Haemoproteus infections from avian blood. J Parasitol. 90(1): 191–194. Doi: 10.1645/GE-3221RN

Xuereb A, Row JR, Brooks RJ, et al. (2012) Relation between Parasitism, Stress, and Fitness Correlates of the Eastern Foxsnake (Pantherophis gloydi) in Ontario. J Herpetol 46:555–561. doi: 10.1670/10-259

1.8 Appendix Online Resources legends

Online Resource 1 Characteristics of localities and reptile species sampled, in the present study.

Online Resource 2 Geographic origin; GenBank lineages and hosts of the haemosporidian parasites included in the phylogenetic analyses.

Online Resource 3 A Bayesian phylogenetic hypothesis of lizard haemosporidian parasites based on partial sequences of cyt b gene (55 sequences and 981 bp excluding gaps). The values above branches are posterior probabilities (see Material and Methods). Sequences from Mammal Plasmodium species were not included in this analysis. 72 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Online Resource 4 Estimates of evolutionary divergence between cyt b sequences of Plasmodium species from lizards. The analysis involved 16 nucleotide sequences and 987 positions in the final dataset. The genetic distances calculated using the Kimura 2- parameter model (Kimura 1980) are shown below the diagonal and, the lowest values are shown in bold. Their standard error estimate(s) are also shown above the diagonal, those were calculated by bootstrap with 500 pseudo-replicates. The rate variation among sites was modeled with a gamma distribution (shape parameter = 4). Online Resource 1 Characteristics of localities and reptile species sampled, in the present study.

Number Locality Genus/ Species of Coordinates Characteristics species Gonatodes 1 riveroi 02º 31’ 44.5” N It presents typical vegetation of Cerro Azul Ninia atrata 1 72º 51’ 59.1” W Guayanese formation, humid tropical 328 m a.s.l forest, and low human intervention. Atractus sp. 1 It is mainly secondary forest of the Ameiva ameiva 1 humid environment; rocks are observed in a state of erosion by humidity and Anolis aff. 1 02º 27’ 56.9” N colonization of some algae and lichens. Ciudad de Piedra auratus 72º 42’ 29.1” W There is an intervened land where the 430 m a.s.l forests have been replaced by pastures. It can be considered as an intervened Mabuya sp. 1 savanna, with outcrops and secondary forest. Cercosaura 1 argulus Leposoma 1 percarinatum Anolis 1 fuscoauratus

Pseudoboa 02º 34’ 18.5” N It is a tropical humid forests and gallery Playa Güio 1 coronata 072º 42’ 49” W forests. 188 m a.s.l Oxyrhopus 1 petolarius Epicrates 1 cenchria Gonatodes 2 riveroi It is rocky outcrops eroded by wind and Anolis auratus 1 2º 32’ 24.5” N water, It has herbaceous and shrub Puentes Naturales 72º 42’51.2” W vegetation at low altitudes, dry climate Plica medemi 2 259 m a.s.l with significant temperature changes between day and night. Leposoma 2 percarinatum Cnemidophorus It is a transition zone between the 2 02º 30’ 49.3” N lemniscatus Orinoquia and the Amazon region, with Puerta de Orión 72º 42’19.7” W a predominance of the rocky substrate 272 m a.s.l Anolis auratus 1 and dry climate.

Anolis scypheus 1 Lepidoblepharis 2 sp. Gonatodes 2 riveroi Helicops 1 angulatus Imantodes 1 cenchoa Mastigodryas 1 boddaerti Corallus 1 hortulanus

Online Resource 2 Geographic origin; GenBank lineages and hosts of the haemosporidian parasites included in the phylogenetic analyses.

GenBank Haemosporidian No. Vertebrate Hosts Geographic origin code Parasites Leucocytozoon 1 AB302215 Gallus gallus domesticus/Chicken Japan caulleryi Leucocytozoon Zonotrichia leucophrys 2 FJ168563 No information majoris oriantha/White-crowned Sparrow Leucocytozoon Pipilo chlorurus/Green-tailed 3 FJ168564 No information fringillinarum towhee Leucocytozoon 4 AB299369 Gallus gallus domesticus/Chicken Malaysia sabrazesi Haemoproteus 5 NC_012448 Columba livia/Rock dove No information columbae Haemoproteus Zenaida galapagoensis/Galapagos 6 KY653756 Galápagos Islands multipigmentatus dove Haemoproteus Creagrus furcatus/Swallow-tailed 7 KY653758 Galápagos Islands jenniae gull 8 KY653777 Haemoproteus iwa Fregata minor/Great frigatebird Galápagos Islands Haemoproteus Dendrocygna autumnalis/Black- 9 KJ499987 Colombia macrovacuolatus bellied whistling duck Haemoproteus Zonotrichia capensis/Rufous- 10 KT698209 Colombia erythrogravidus collared sparrow Haemoproteus Arremon brunneinucha/Chestnut- 11 KT698210 Colombia coatneyi capped brush finch Eriocnemis derbyi/Black-thighed 12 KY653794 Haemoproteus witii Colombia puffleg Haemoproteus 13 KY653799 Luscinia luscinia/Thrush nightingale Lithuania balmorali 14 KY653787 Haemoproteus lanii Lanius collurio/Red-backed shrike Lithuania Haemoproteus 15 KY653790 Hippolais icterina/Icterine warbler Lithuania belopolskyi Haemoproteus 16 KY653793 Sturnus vulgaris/Common starling Lithuania pastoris Haemoproteus 17 KY653763 Turdus merula/Common blackbird Lithuania minutus Haemoproteus Motacilla flava/Western yellow 18 KY653768 Lithuania motacillae wagtail Haemoproteus 19 KY653807 Loxia curvirostra/Red crossbill Lithuania tartakovskyi Haemoproteus 20 KY653757 Asio otus/Long-eared owl Lithuania noctuae Haemoproteus 21 KY653811 Zenaida macroura/Mourning dove Arizona-USA sacharovi Haemocystidium Teratoscincus scincu/Frog-eyed 22 AY099062 Pakistan kopki gecko Haemocystidium Ptyodactylus hasselquisti/Fan-footed 23 AY099057 Israel ptyodactylii gecko Haemocystidium 24 KF049514 Naja annulifera/Snouted cobra South Africa mesnili Haemocystidium Testudo graeca/Spur-thighed 25 KM068153 Iran anatolicum tortoise Haemocystidium Podocnemis unifilis/Yellow-spotted 26 KF049506 Peru pacayae river turtle 27 KF049492 Haemocystidium Podocnemis expansa/Arrau turtle Peru peltocephali Plasmodium Sceloporus occidentalis/Western 28 NC_009960 (Paraplasmodium) No information fence lizard mexicanum Plasmodium 29 NC_009961 (Lacertamoeba) Anolis sagrei/Brown anole No information floridense Plasmodium MF177708 30 (Sauramoeba) Cnemidophorus gramivagus Colombia

kentropyxi MF177707 Plasmodium 31 Plica cf. plica Colombia (Lacertamoeba) sp. Plasmodium 32 MF177709 (Carinamoeba) Ameiva ameiva/Giant ameiva Colombia carmelinoi Plasmodium Sceloporus jarrovii/Yarrow's Spiny 33 KY653779 (Paraplasmodium) Arizona-USA Lizard chiricahuae 34 AY099048 Plasmodium agamae Agama agama/Common agama Ghana/Africa Plasmodium 35 AY099053 Agama agama/Common agama Ghana/Africa giganteum Plasmodium Caribbean island of 36 KR477594 No information hispaniolae Hispaniola Plasmodium Caribbean island of 37 KR477583 No information fairchildi Hispaniola Plasmodium 38 AY099055 Anolis oculatus/Dominican anole Dominica Island azurophilum Plasmodium Emoia longicauda/Shrub Whiptail- 39 EU834710 Papua New Guinea lacertiliae skink Prasinohaema Plasmodium 40 EU834703 prehensicauda/Prehensile Green Papua New Guinea minuoviride Tree Skink Plasmodium 41 EU834705 Sphenomorphus simus Papua New Guinea megalotrypa Plasmodium 42 EU834704 Sphenomorphus jobiensis Papua New Guinea koreafense Hypsilurus modestus/Modest Forest 43 EU834707 Plasmodium gemini Papua New Guinea Dragon Plasmodium Pseudocordylus melanotus/Highveld 44 KX121607 South Africa intabazwe Crag Lizard Plasmodium Cordylus vittifer/Transvaal Girdled 45 KX121609 South Africa zonuriae Lizard Plasmodium 46 AB250690 Gallus gallus domesticus/Chicken Philippines gallinaceum Plasmodium 47 NC_008279 Gallus gallus domesticus/Chicken Japan juxtanucleare 48 KC138226 Plasmodium lutzi Turdus fuscater/Great thrush Colombia 49 KY653814 Plasmodium unalis Turdus fuscater/Great thrush Colombia Plasmodium Zonotrichia capensis/Rufous- 50 KY653770 Colombia homopolare collared sparrow Plasmodium 51 KY653775 Sylvia borin/Garden warbler Lithuania ashfordi Plasmodium 52 KY653792 Turdus merula/Common blackbird Lithuania vaughani Plasmodium 53 KY653772 Passer domesticus/House sparrow Lithuania relictum Plasmodium 54 KY653784 Lanius collurio/Red-backed shrike Lithuania homocircumflexum Plasmodium Troglodytes troglodytes/Eurasian 55 KY653762 Lithuania circumflexum wren Plasmodium Acrocephalus scirpaceus/Eurasian 56 KY653801 Lithuania elongatum reed warbler Plasmodium Alopochen aegyptiacus/Egyptian 58 JX467689 Brazil nucleophilum goose 59 AY598140 P. vivax Homo sapiens Salvador 60 AB444129 P. cynomolgi Macaca nemestrina Malaysia Macaca sinica, M. nemestrina, M. 61 GQ355483 P. inui South and East Asia fascicularis, M. mulatta 62 AB354573 P. hylobati Hylobati moloch Indonesia, Malaysia M. nemestrina, M. fascicularis, M. 63 NC_007232 P. knowlesi Southeast Asia nigra M. radiata, M. mulatta, Prebytis Southern India, Sri 64 AY722799 P. fragile spp. Lanka 65 AY800111 P. gonderi Cercocebus atys, Cercopithecus spp. Central Africa Tropical, subtropical, 66 AB354570 P. malariae Homo sapiens and temperate regions Tropical and 67 HQ712053 P. ovale-wallikeri Homo sapiens subtropical regions Madagascar (Eastern 68 HQ712054 Plasmodium sp. (A) Hapalemur griseus griseus rainforest) Madagascar (Eastern 69 HQ712055 Plasmodium sp. (B) Varecia variegata rainforest) 70 AF014115 P. berghei Grammomys sp. Central Africa 71 AF014116 P. chabaudi Thamnomys sp. Central Africa 72 NC-002235 P. reichenowi Pan troglodytes Africa Worldwide Tropical 73 AY282930 P. falciparum Homo sapiens regions Thailand: Mueang 74 LC090213 Plasmodium bubalis Bubalus bubalis Mukdahan 75 LC090215 Plasmodium sp. Capra aegagrus hircus Zambia: Chama

Online Resource 1 A Bayesian phylogenetic hypothesis of lizard haemosporidian parasites based on partial sequences of cyt b gene (55 sequences and 981 bp excluding gaps). The values above branches are posterior probabilities (see Material and Methods). Sequences from Mammal Plasmodium species were not included in this analysis.

Online Resource 4 Estimates of evolutionary divergence between cyt b sequences of Plasmodium species from lizards. The analysis involved 16 nucleotide sequences and 987 positions in the final dataset. The genetic distances calculated using the Kimura 2-parameter model (Kimura 1980) are shown below the diagonal and, the lowest values are shown in bold. Their standard error estimate(s) are also shown above the diagonal, those were calculated by bootstrap with 500 pseudo-replicates. The rate variation among sites was modeled with a gamma distribution (shape parameter = 4).

Plasmodium species 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16

1 P. agamae (AY099048) 0.007 0.007 0.007 0.007 0.008 0.007 0.008 0.007 0.009 0.010 0.009 0.008 0.010 0.008 0.009 2 P. giganteum (AY099053) 0.039 0.007 0.007 0.007 0.008 0.007 0.008 0.007 0.009 0.011 0.009 0.007 0.011 0.007 0.010

3 P. azurophilum (AY099055) 0.051 0.044 0.005 0.006 0.007 0.006 0.006 0.004 0.008 0.010 0.008 0.006 0.010 0.006 0.008

4 P. minuoviride (EU834703) 0.050 0.048 0.028 0.004 0.006 0.006 0.007 0.006 0.008 0.010 0.008 0.006 0.010 0.007 0.008

5 P. koreafense (EU834704) 0.050 0.048 0.038 0.019 0.006 0.007 0.007 0.006 0.009 0.011 0.008 0.007 0.011 0.007 0.009

6 P. megalotrypa (EU834705) 0.059 0.061 0.042 0.030 0.032 0.007 0.008 0.007 0.009 0.010 0.009 0.007 0.011 0.008 0.009

7 P. gemini (EU834707) 0.052 0.051 0.039 0.040 0.043 0.049 0.007 0.006 0.008 0.010 0.009 0.007 0.010 0.007 0.009

8 P. lacertiliae (EU834710) 0.063 0.059 0.036 0.038 0.045 0.049 0.054 0.006 0.009 0.011 0.010 0.007 0.011 0.008 0.010

9 P. fairchildi (KR477583) 0.052 0.046 0.015 0.029 0.033 0.039 0.045 0.031 0.008 0.009 0.008 0.006 0.010 0.006 0.008

10 P. hispaniolae (KR477594) 0.074 0.075 0.058 0.058 0.064 0.065 0.058 0.067 0.063 0.011 0.008 0.009 0.012 0.009 0.008

11 P. mexicanum (NC_009960) 0.096 0.095 0.086 0.085 0.093 0.094 0.088 0.094 0.083 0.102 0.010 0.010 0.005 0.010 0.011

12 P. floridense (NC_009961) 0.069 0.071 0.060 0.058 0.065 0.072 0.068 0.072 0.060 0.057 0.092 0.008 0.011 0.008 0.004

13 P. carmelinoi (MF177709) 0.055 0.053 0.037 0.044 0.045 0.052 0.052 0.050 0.035 0.073 0.100 0.068 0.011 0.005 0.009

14 P. chiricahua (KY653779) 0.100 0.101 0.092 0.091 0.101 0.100 0.092 0.096 0.088 0.114 0.024 0.102 0.106 0.011 0.011

15 P. kentropyxi (MF177708) 0.061 0.055 0.042 0.050 0.050 0.061 0.055 0.052 0.037 0.081 0.101 0.072 0.020 0.107 0.009

16 P. (Lacertamoeba) sp. (MF177707) 0.078 0.078 0.067 0.066 0.068 0.073 0.072 0.080 0.067 0.058 0.093 0.017 0.073 0.104 0.076

IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

2. Capítulo 2. Haemocystidium spp., a species complex infecting ancient aquatic turtles of the family Podocnemididae: First report of these parasites in Podocnemis vogli from the Orinoquia Haemocystidium spp., Un complejo de especies que infecta a las tortugas acuáticas antiguas de la familia Podocnemididae: primer informe de estos parásitos en Podocnemis vogli de la Orinoquia

El género Haemocystidium fue descrito en 1904 por Castellani y Willey. Sin embargo, varios estudios lo consideraron un sinónimo de los géneros Plasmodium o Haemoproteus. Recientemente, la evidencia molecular ha demostrado la existencia de un grupo monofilético que corresponde al género Haemocystidium. Aquí, exploramos más a fondo el clado Haemocystidium sp. mediante el estudio de los parásitos de Testudinos. Se analizaron un total de 193 individuos pertenecientes a seis familias de Testudinos. Las muestras fueron recolectadas en cinco localidades de Colombia: Casanare, Vichada, Arauca, Antioquia y Córdoba. De cada individuo, se tomó una muestra de sangre para análisis moleculares y se realizaron frotis de sangre periférica, que se fijaron y se tiñeron con Giemsa. La prevalencia de Haemocystidium spp. fue 1.55% (n = 3/193); todos los individuos infectados pertenecían a Podocnemis vogli (Savanna Side-necked Turtle) del departamento de Vichada. Este es el primer reporte de Haemocystidium spp. En Colombia e infectando esta especie. El análisis filogenético de un fragmento mitocondrial reveló Haemocystidium spp. como grupo monofilético y como taxón hermano de Haemoproteus catharti y del género Plasmodium. El parásito encontrado en nuestro estudio es morfológicamente indistinguible de Haemocystidium (Simondia) pacayae reportado en Perú y Brasil. Sin embargo, un nuevo linaje encontrado en P. vogli muestra una distancia genética de 0.02 con Hae. pacayae y 0.04 con Hae. peltocephali. Se propone que este 81 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA. resultado debe ser complementado con mas información para la descripción de especies. Haemocystidium spp. son difíciles de identificar solo por morfología. Como resultado, es posible que algunos de los taxones, como Haemocystidium (Simondia) pacayae, sea un complejo de especies crípticas.

Palabras claves: Hemoparásito, reptil, Simondia, Quelonios, Colombia.

Este artículo abarca los tres objetivos del proyecto de maestría, donde se reporta la presencia de Haemocystidium sp, con base en la descripción morfológica de las formas observadas en sangre periférica y el análisis molecular del fragmento de citocromo b. Mi contribución a este artículo radicó en la revisión de los extendidos sanguíneos de los individuos incluidos en el estudio, la toma de fotografías y medidas, así como la identificación morfológica de la especie, ademas el análisis detallado de la morfología de cada uno de los morfotipos, determinar las parasitemias y prevalencia. También, contribuí al análisis de las secuencias obtenidas, y la escritura de todo el artículo. Esta es la versión pre-print. Este manuscrito fue aceptado en International Journal of Parasitology: Parasites and Wildlife

IJP: Parasites and Wildlife 10 (2019) 299–309

Contents lists available at ScienceDirect

IJP: Parasites and Wildlife

journal homepage: www.elsevier.com/locate/ijppaw

Haemocystidium spp., a species complex infecting ancient aquatic turtles of the family Podocnemididae: First report of these parasites in Podocnemis T vogli from the Orinoquia

Leydy P. Gonzáleza,b, M. Andreína Pachecoc, Ananías A. Escalantec, Andrés David Jiménez Maldonadoa,d, Axl S. Cepedaa, Oscar A. Rodríguez-Fandiñoe, ∗ Mario Vargas‐Ramírezd, Nubia E. Mattaa, a Departamento de Biología, Facultad de Ciencias, Universidad Nacional de Colombia, Sede Bogotá, Carrera 30 No 45-03, Bogotá, Colombia b Instituto de Biotecnología, Facultad de Ciencias, Universidad Nacional de Colombia, Sede Bogotá, Carrera 30 No 45-03, Bogotá, Colombia c Department of Biology/Institute for Genomics and Evolutionary Medicine (iGEM), Temple University, Philadelphia, PA, USA d Instituto de Genética, Universidad Nacional de Colombia, Sede Bogotá, Carrera 30 No 45-03, Bogotá, Colombia e Fundación Universitaria-Unitrópico, Dirección de Investigación, Grupo de Investigación en Ciencias Biológicas de la Orinoquía (GINBIO), Colombia

ARTICLE INFO ABSTRACT

Keywords: The genus Haemocystidium was described in 1904 by Castellani and Willey. However, several studies considered Haemoparasites it a synonym of the genera Plasmodium or Haemoproteus. Recently, molecular evidence has shown the existence Reptile of a monophyletic group that corresponds to the genus Haemocystidium. Here, we further explore the clade Simondia Haemocystidium spp. by studying parasites from Testudines. A total of 193 individuals belonging to six families of Chelonians Testudines were analyzed. The samples were collected in five localities in Colombia: Casanare, Vichada, Arauca, Colombia Antioquia, and Córdoba. From each individual, a blood sample was taken for molecular analysis, and peripheral blood smears were made, which were fixed and subsequently stained with Giemsa. The prevalence of Haemocystidium spp. was 1.55% (n = 3/193); all infected individuals belonged to Podocnemis vogli (Savanna Side-necked turtle) from the department of Vichada. This is the first report of Haemocystidium spp. in Colombia and in this turtle species. The phylogenetic analysis of a mitochondrial cytb fragment revealed Haemocystidium spp. as a monophyletic group and as a sister taxon of Haemoproteus catharti and the genus Plasmodium. Haemocystidium spp. are difficult to identify by morphology only. As a result, it is possible that some of the taxa, such as Haemocystidium (Simondia) pacayae, represent a species complex. The parasite found in our study is morphologically indistinguishable from Haemocystidium (Simondia) pacayae reported in Peru. However, the new lineage found in P. vogli shows a genetic distance of 0.02 with Hae. pacayae and 0.04 with Hae. peltocephali.Itis proposed that this divergent lineage might be a new species. Nevertheless, additional molecular markers and ecological features could support this hypothesis in the future.

1. Introduction cause human malaria (WHO, 2018), the information for other taxonomic groups that parasitize wildlife is limited and fragmented. Such a The phylum Apicomplexa (Levine, 1988) encompasses at least knowledge gap has driven several phylogenetic studies within the order 5,000 recognized species of obligate intracellular protozoan parasites Haemosporida (Javanbakht et al., 2015; Maia el at., 2016; Boundenga (Morrison, 2009). Among them, blood parasites belonging to the order et al., 2017; Pacheco et al., 2018b). The lack of information is parti- Haemosporida (Danilewsky 1885) are classified into the families Leu- cularly critical in haemosporidian parasites in reptiles. cocytozoidae (infecting birds), Haemoproteidae, and Garniidae (found The haemosporidians found in reptiles have been classified into in birds and reptiles), and Plasmodiidae (which infects birds, reptiles, several families; Haemoproteidae, genera Haemoproteus sp. and and mammals). Despite significant advances in the description of spe- Haemocystidium sp.; Plasmodidae, genus Plasmodium sp.; Garnidae, cies diversity of the genus Plasmodium, which includes the species that genera Garnia sp. and Fallisia sp. (Levine, 1970; Adl et al., 2012, 2019;

Abbreviations:H: Haemoproteus, Hae: Haemocystidium ∗ Corresponding author. E-mail address: [email protected] (N.E. Matta). https://doi.org/10.1016/j.ijppaw.2019.10.003 Received 26 June 2019; Received in revised form 17 October 2019; Accepted 21 October 2019 2213-2244/ © 2019 The Authors. Published by Elsevier Ltd on behalf of Australian Society for Parasitology. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/BY-NC-ND/4.0/). L.P. González, et al. IJP: Parasites and Wildlife 10 (2019) 299–309

Fig. 1. Timeline of the taxonomic classiﬁcation of the genus Haemocystidium. Relevant events in the description of this genus.

Perkins, 2014); and the family Leucocytozoidae, genus Saurocytozoon (Castaño-Mora, 2002; Rueda-Almonacid et al., 2007). Specimens from sp. (Lainson et al., 1974). Among those, the genus Haemocystidium has three species of the genus Podocnemis from the Podocnemididae family been one of the least studied, and its status has been controversial. were analyzed. This was a diverse group in the Late Cretaceous (at least Being synonymized as Plasmodium or Haemoproteus, after its description 20 genera and 30 species) that was reduced to only three extant genera (Castellani, and Willey, 1904, Fig. 1). (Podocnemis, Peltocephalus, and Erymnochelys) and eight species Currently, the genus Haemocystidium has not been fully accepted by (Gaffney et al., 2011) distributed in South America and Madagascar the academic community due to the limited studies available in this (Vargas-Ramirez et al., 2008). In this study, Podocnemis vogli, P. le- group, both in terms of hosts and parasite species (Javanbakht et al., wyana, and P. unifilis were screened for haemoparasites. Podocnemis 2015). New molecular lineages attributed to this genus have been re- vogli (Savanna Side-necked Turtle) is an omnivorous and aquatic spe- ported in samples from the gecko species Hemidactylus luqueorum and cies, which restricts its distribution to the Orinoco basin in the east of Ptyodactylus hasselquistii captured in Oman, Asia (Maia et al., 2016), Colombia and western plains of Venezuela (Rueda-Almonacid et al., and in turtles, Pelusios castaneus and Kinixys erosa, from Gabon, Central 2007). Podocnemis unifilis is distributed in Colombia, Venezuela, Brazil, Africa (Supplementary Table 1; Boundenga et al., 2017). The few re- Peru, Ecuador, Bolivia, and the Guianas, inhabiting white and black ports from South America have found three Haemocystidium species: waters (Rueda-Almonacid et al., 2007). Podocnemis lewyana, is also a Haemocystidium (S.) pacayae infecting the freshwater turtles Podocnemis Colombian endemic species with a restricted distribution to the basins unifilis and Podocnemis expansa (Peru), Hae. geochelonis in the tortoise of the Magdalena and Sinú rivers; it is the only species of this family Chelonoidis denticulata (Brazil) and Hae. (S.) pelthocephali in Peltoce- that inhabits the west of the Andes mountain range, and it has diurnal phalus dumerilianus, Podocnemis unifilis, and P. expansa (Brazil and and nocturnal habits (Rueda-Almonacid et al., 2007). From the Geoe- Peru), (Lainson and Naiff, 1998; Pineda-Catalan et al., 2013). However, mydidae family, Rhinoclemmys melanosterna is distributed in the Pacific, there is an extraordinary diversity of turtles in Colombia, and there are Caribbean region and lower Magdalena river (Rueda-Almonacid et al., no studies directed to assess the presence and diversity of haemospor- 2007); it preferably inhabits swampy areas where heliconias are idian infecting these hosts. abundant. Belonging to the Testudinidae family, Chelonoidis carbonaria There are nine families and 36 species of turtles in Colombia, 28 are is a terrestrial tortoise with a broad distribution that includes Vene- freshwater turtles, two are exclusively terrestrial, and six are sea turtles. zuela, Colombia, the Guyanas, Brazil, Bolivia, Paraguay, and Argentina Fifteen of these species are classified into various categories of threat (Castaño-Mora, 2002; Rueda-Almonacid et al., 2007). Belonging to the (Ministerio del Medio Ambiente, Vivienda y Desarrollo Territorial, Kinosternidae family, Kinosternon scorpioides is distributed from Mexico 2017). The few studies conducted on Testudines have focused on their to Argentina, inhabiting rivers, lakes, and flood plains (Rueda- microorganisms such as bacteria (Pachón, 2009) or pathological studies Almonacid et al., 2007). of some of their parasites such as Toxoplasma spp. and Filaroides spp. All sampled chelonians were captured by using funnel traps, fishing (Herrera, 2008), while the biodiversity of haemoparasites that may nets, and by hand. The blood samples were obtained from the sub- affect them is poorly known. This study had two objectives: (1) To carapacial sinus or the coccygeal vein and did not exceed 1% of the analyze the presence of haemosporidian parasites in wild turtles be- body weight. For each individual, three peripheral blood smears were longing to six families distributed in diverse ecoregions in Colombia; made, which were dried immediately and fixed with absolute methanol and (2) to discuss the classification of species belonging to Haemocys- for 5 min. At the laboratory, the blood smears were stained with Giemsa tidium genus based on morphological and molecular data. (4%, pH 7.2) for 45 min. The blood samples were preserved in absolute ethanol and stored at −20 °C for further molecular analysis. The sam- fi − 2. Materials and methods ples were kept at room temperature in the eld and then at 20 °C in the laboratory. 2.1. Sampling

In total, 193 wild turtles (order Testudines) belonging to nine spe- 2.2. Ethical statement and sampling permits cies representing the families Chelidae, Podocnemididae, Kinosternidae, Emydidae, Geoemydidae, and Testudinidae were sam- Specimens were collected under the framework collection permis- pled. These individuals were collected in the departments of Antioquia, sion for wild species of biological diversity authorized by the National Arauca, Casanare, Cordoba, and Vichada (Fig. 2). The numbers of in- Environmental Licenses Authority (ANLA) to the Universidad Nacional dividuals sampled by species and their corresponding localities are re- de Colombia by resolution No. 0255 of March 14, 2014. All turtles and ported in Table 1. Below there is a brief description and distribution of tortoises were released after the blood samples were collected. The “ the species included in this study. From Chelidae family, Mesoclemmys samples analyzed in this study came from the collection Banco de te- ” dahli is a Colombian trans-Andean endemic species that, except for the jidos de la Biodiversidad Colombiana, Universidad Nacional de other species belonging to this family (genera Chelus, Platemys, Rhin- Colombia. emys, and Phrynops), is distributed in the tropical dry forest along the Caribbean, inhabiting shallow waters with ﬂoating vegetation

300 L.P. González, et al. IJP: Parasites and Wildlife 10 (2019) 299–309

Fig. 2. Species and sampling locations for turtles analyzed in the study. Turtle species present in these localities are shown.

2.3. Blood ﬁlm examination genomic DNA template, 12.5 μL of DreamTaq Master Mix (Thermo Fisher Scientiﬁc, Germany), 8.5 μL nuclease-free water, and 1 μL of each Blood slides were examined using an Olympus CX41 microscope primer. In the case of the nested PCR, reactions were performed in (Olympus Corporation, Tokyo, Japan), at 400× for 10 min, and then at 50 μL total volume, with 3 μL of the primary PCR products, 1× PCR

1000× for 20 min. Slides with haemoparasites were examined entirely, buffer, 2.5 mM MgCl2, 0.3 units of Taq DNA polymerase (Thermo Sci- and digital images were obtained using an Olympus DP27 digital entific, Waltham, USA), 0.2 mm dNTPs (Promega, Madison, USA), and camera and processed with the CellSens software (Olympus 0.4 mM of each primer. One negative control (nuclease-free water) and Corporation, Tokyo, Japan). Morphometric analysis was performed one positive control (Plasmodium unalis positive sample) were included using ImageJ software (Schneider et al., 2012), and the parasitaemia in each PCR. Temperature profiles were the same as in the original (No. of parasites/10,000 erythrocytes) was estimated from erythrocyte protocol (Supplementary Table S2; Pacheco et al., 2018a). All products counts at a magnification of 1000× on areas where blood cells formed from the primary and nested PCR were evaluated by running 2 μL of the a monolayer (Staats and Schall, 1996). The taxonomic determination of final products on 2% agarose gels. Amplified products were cleaned haemoparasites was made comparing their morphologies and mor- using differential precipitation with ammonium acetate protocol phometry with previous reports in the literature for Haemoproteus and (Bensch et al., 2000). Fragments of DNA from all positive amplifications Haemocystidium species (Telford, 2009; Lainson, 2012; Pineda-Catalan were sequenced in both directions using a 3730xl DNA Analyzer (Ap- et al., 2013; Maia et al., 2016). The comparison of morphological plied Biosystems, Foster City, CA) through Macrogen (Macrogen Inc.). measurements between the Haemocystidium species found here and The sequences for the cytb fragments obtained in this study were those previously reported was done using the Student's t-test. identified as Haemocystidium species using BLAST (Altschul et al., 1997). The sequences were deposited in GenBank under the accession numbers MK976708 (GERPH: PC004), MK976709 (GERPH: PC006) 2.4. DNA extraction and detection of haemosporidian parasites by a and MK976710 (GERPH: PC005). polymerase chain reaction

Total DNA was extracted from samples using the phenol-chloroform 2.5. Phylogenetic analysis of the parasite cytb gene fragment method (Sambrook et al., 1989) and measured by using NanoDrop Lite spectrophotometer (Thermo Scientific, Massachusetts, USA). A nested In order to link morphological characteristics with molecular polymerase chain reaction (PCR) assay using primers targeting the lineages, only positive samples by microscopy were used in this in- parasite cytochrome b (cytb) gene from the mitochondria was used vestigation. Of those, one sample amplified in the primary PCR (Supplementary Table S2; Pacheco et al., 2018a). Reactions for primary (GERPH: PC005) described above, and two in the nested PCR (GERPH: PCR were performed in 25 μL total volume, including 2 μL of total PC004 and GERPH: PC005), thus two different fragment sizes for the

301 L.P. González, et al. IJP: Parasites and Wildlife 10 (2019) 299–309

cytb gene were obtained in this study (1,692 and 535 bp without the primer regions respectively). In order to compare our molecular data 134 2 1 against all sequences that have been published for Haemocystidium and ″ ″ other haemoparasites infecting reptiles or birds (Plasmodium, W 3*/13 . W1

W5 ﬀ W4 ″ Haemoproteus and Leucocytozoon), two di erent nucleotide alignments ″ ″ 47.5309 47.5309 ″ ′ ′ were performed using ClustalX v2.0.12 and Muscle as implemented in 742 ′ 31.05

19.69 ﬁ

19.69 SeaView v4.3.5 (Gouy et al., 2010) with manual editing. The rst ′ ′ ′ N 71° 14 N 71° 14 alignment was constructed with 103 cytb partial sequences (233 bp ″ ″ N 67°49 excluding gaps) belonging to four genera (Leucocytozoon, Haemoproteus, ″ Haemocystidium N 70° 14 N 74° 12 N. 74° 12

″ Haemocystidium, and Plasmodium). Although our smallest cytb fragment W 067 ″ ″ ′ ″ 18.9463 18.9463 ′ ′ for samples GERPH: PC004 and GERPH: PC006 was 535 bp in length, 30.84

′ the region that overlapped between all sequences, including those 48.311 27.0720 27.0720 ′ ′ ′ available in the databases, was only 233 bp in length. This alignment included the sequences obtained in this study for samples GERPH: PC004, GERPH: PC005 and GERPH: PC006, as well as sequences from N. 69° 30

″ well-known parasite species based on morphology and haplotypes (Valkiūnas and Iezhova, 2018) that were available on the GenBank

26.796 database (Benson et al., 2012) at the time of this study. The second ′ alignment was constructed with 62 sequences corresponding to a bigger Puerto Carreño (Vichada) 06°16 W * Lorica (Cordoba)Lorica (Cordoba)Yondó (Antioquia) 6° 48 Yondó (Antioquia) 6° 48 3 11 Cumaribo (Vichada) 3° 22 W Paz de Ariporo (Casanare) 5° 42 Paz de Ariporo (Casanare) 5° 42 Lorica (Cordoba) 17 Sample locality (Department)Loríca (Cordoba) n 3 fragment of cytb (707 bp excluding gaps) also belonging to the four genera, including only the sequence obtained for GERPH: PC005 and sequences from well-known parasite species (using morphology) available on GenBank. In this case, our larger cytb fragment obtained

The only locality that has turtles infected with for GERPH: PC005 had 1,692 bp, but the region that overlapped among * all the sequences included was 707 bp. Phylogenetic trees were inferred based on the first (Supplementary Fig. S1) and second alignments (Fig. 3A), using a Bayesian method as implemented in MrBayes v3.2.6 with the default priors (Ronquist and Huelsenbeck, 2003) and the general time-reversible model with gamma-distributed substitution rates and a proportion of invariant sites (GTR +Γ + I). This model was the one with the lowest Bayesian In- formation Criterion (BIC) scores for both alignments as estimated by MEGA v7.0.14 (Kumar et al., 2016). Posterior probability was estimated for the nodes in MrBayes by sampling every 1,000 generations from two independent chains lasting 3Χ106 Markov Chain Monte Carlo (MCMC) steps. The chains were assumed to have converged when the average SD of the posterior probability was < 0.01 and the value of the potential scale reduction factor (PSRF) was between 1.00 and 1.02 (Ronquist and Huelsenbeck, 2003). Then, 25% of the sample was discarded once convergence was reached as a “burn-in.” For both phylogenies, Leucocytozoon species were used as out-group. Genbank accession numbers for all sequences used in the analyses are given in the phylogenetic trees. Furthermore, in order to estimate the genetic distance between Haemocystidium species, the number of base substitutions per site and the standard error estimates between sequences of species from well- known parasites based on morphology were estimated using the Tamura-Nei model. This model was the one that better fit to the data (Tamura and Nei, 1993). In a second analysis, the number of base substitutions per site and the standard error estimates from averaging overall sequence pairs (haplotypes) within and between species were Colombian slider Colombian Wood Turtle Scorpion Mud turtle Red-footed Tortoise Yellow-footed Tortoise, Brazilian Giant Tortoise, Forestfooted Tortoise, Tortoise South American Tortoise, South American Yellow- Savanna Side-necked Turtle Common name Magdalena River Turtle Dahl's Toad-headed Turtle Yellow-spotted River Turtle, Yellow-headed Side-neck, Yellow-spotted Side-neck Turtle Cravo Norte (Arauca) 06°14 also estimated. Codon positions included were 1st+2nd+3rd, and all positions containing gaps and missing data were eliminated. These analyses were conducted in MEGA7 (Kumar et al., 2016) using the second alignment (707 bp excluding gaps). lis fi 3. Results

3.1. Morphological detection of haemosporidian parasites Trachemys venusta callirostris Rhinoclemmys melanosterna Kinosternon scorpioides Chelonoidis carbonaria Chelonoidis denticulata Podocnemis vogli Species Podocnemis lewyana Mesoclemmys dahli Podocnemis uni Of the 193 captured turtles, only three individuals were positive for haemosporidian parasites using microscopy (3/193, 1.55%), all belonging to the same host species, P. vogli. The only haemosporidian detected in the samples analyzed were Haemocystidium, and its prevalence in P. vogli was 2.04% (n = 3/147). All three positive samples Emydidae Geoemydidae Kinosternidae Testudinidae Family Chelidae Podocnemididae

Table 1 Turtle species analyzed in the present study, from diverse localities in Colombia. Common names, size sample, and locality are shown. came from a single locality: Finca de las Flores in the department of

302 L.P. González, et al. IJP: Parasites and Wildlife 10 (2019) 299–309

Fig. 3. (A) A Bayesian phylogenetic analysis of reptile haemosporidian parasites based on 62 partial sequences of cytb gene sequences corresponding to a bigger fragment of cytb (707 bp excluding gaps). Leucocytozoon genus was used as outgroup. In parenthesis are GenBank sequence accession number, isolate name, and turtle species name respectively. Branch color indicates the parasites genus: Blue, Plasmodium sp.; light green, Haemocystidium spp. infecting lizards and snakes; dark green, Haemocystidium sp. Infecting turtles; black, Haemoproteus and Leucocytozoon spp. (B–C) Estimates of evolutionary divergence between/within Haemocystidium spp. Genetic distances were estimated using the bigger fragment of cytb (707 bp excluding gaps). The number of base substitutions per site between sequences are shown in black and the standard error estimate(s) are shown above the diagonal in blue. Evolutionary divergence between/within Haemocystidium pacayae and Haemocystidium sp. (GERPH: PC005) are shown in bold and red respectively. Sequences previously identiﬁed as Hae. pacayae (KF049495 and KF049507) are likely Hae. (Simondia) sp. (group 2). (For interpretation of the references to color in this ﬁgure legend, the reader is referred to the Web version of this article.)

Fig. 4. Haemocystidium (Simondia) sp. (GERPH: PC005) identified in Podocnemis vogli (A–H). (A–F) Young gametocytes, (E) coinfection with a gamont of Haemogregarina, (G) microgametocyte, and (H) macrogametocyte. Haemocystidium (S.) pacayae (GERPH: PC004-PC006) identified in P. vogli (I–J). (I) Young gametocyte, (K) microgametocyte, and (L) macrogametocyte. Bold black arrow: hemozoin granules; white arrow: parasitophorous vacuole; fine black arrow: vacuole. Giemsa stain, Scale bar: 10 μm.

303 L.P. González, et al. IJP: Parasites and Wildlife 10 (2019) 299–309

Vichada, where 13 P. vogli were captured (06°16.067′ N. 67°49.742′ W) the vacuole in some instances is big, so the morphology of the game- so the prevalence of Haemocystidium sp. in this area was 23.1% (Fig. 2, tocyte resembles the ring stage typical in Plasmodium falciparum Table 1). Parasites were found infecting mature erythrocytes (Fig. 4). (Fig. 4E). In at least 12.5% of the cases, it was possible to observe a The parasitaemia was 0.35% for the individual GERPH: PC004, 0.68% structure similar to a parasitophorous vacuole (PV) described in other for GERPH: PC005, and 0.29% for GERPH: PC006. The most frequent haemoparasites. In these cases, the parasite and PV together measured haemoparasites observed in these samples were haemogregarines between 5.87 and 30.12 μm2 (Fig. 4F). (84.53%, n = 164/193). All individuals infected with Haemocystidium showed co-infection with haemogregarine gamonts (data not shown). 3.3.2. Microgametocytes All the slides were deposited in the biological collection of “Grupo de Microgametocytes are rounded (9.39 × 5.88 μm), the pigment Estudio Relación Parásito Hospedero (GERPH)” at the Department of granules are small, and distributed both randomly and at the poles of Biology, Universidad Nacional de Colombia, Bogotá, Colombia. the parasite (Fig. 4G). The parasite nucleus is diﬀuse, located at the periphery. Occasionally, small vacuoles can be observed in the parasite 3.2. Genetic distances and phylogenetic relationships of Haemocystidium (Fig. 4G). The proportion of macrogametocytes:microgametocytes was genus 1:2.

A fragment of cytb gene was obtained for each infected turtle. The 3.3.3. Macrogametocytes genetic distances between Haemocystidium species estimated as the Cytoplasm is granular in appearance and pigment granules may number of base substitutions per site are shown in Fig. 3B. The cytb form clumps (Fig. 4H). Macrogametocytes might have small vacuoles gene fragments (535 bp) of samples GERPH: PC004 and GERPH: PC006 (Fig. 4H), but in advanced stages, vacuoles are conspicuous (Fig. 4H). were 100% identical to the sequence reported for Haemocystidium pa- Nucleus is compact and located laterally. We did not find significant cayae (KF049506; Pineda-Catalan et al., 2013; Supplementary Fig. S1). differences (p < 0.001) at the morphometric level between Haemo- However, the genetic distance between Haemocystidium pacayae cystidium sp., and Hae. (S.) pacayae, and Hae. (S.) peltocephali (Table 2). (KF049506, KF049509, and KF049513) and the parasite infecting However, the genetic distance of the cytb fragments between Hae. (Si- sample GERPH: PC005 was 0.02 ± 0.006 (Fig. 3B). mondia) sp. (GERPH: PC005) and Hae. pacayae (e. g., KF049506) is Bayesian phylogenetic trees of Haemocystidium parasites, generated 0.02, and Hae. (Simondia) sp. with Hae. (S.) peltocephali (KF049492) is using the first (103 cytb partial sequences with 233 bp; Supplementary 0.04 (Fig. 3B and C). Fig. S1) and the second alignments (62 cytb partial sequences with 707 bp; Fig. 3A) showed similar topologies. All the species identified as 4. Discussion Haemocystidium parasites infecting reptiles (snakes, lizards, and turtles) form a monophyletic group sharing a common ancestor with Haemo- The reports of Haemocystidium sp. worldwide are limited. Thus this proteus catharti and the genus Plasmodium. Two lineages of Haemocys- study represents the third report of species belonging to this genus in tidium parasites identified as 1 and 2 (Fig. 3C), were found infecting South America and the first in Colombian Podocnemidids. It is im- three individuals of P. vogli and were grouped with the lineages re- portant to highlight that the overall prevalence of Haemocystidium ported so far as Hae. pacayae. The lineage found infecting the samples found in P. vogli from Colombia (2.04%, 3/147) is lower than that GERPH: PC004 and GERPH: PC006 (group 1) was identical to the Hae. previously reported by Pineda-Catalan et al. (2013) in P. expansa of pacayae sequences available on the GenBank database under the ac- 12.5% (12/96) and P. unifilis of 9.5% (13/136). These differences could cession numbers KF049506, KF049509, and KF049513 (Pineda-Catalan be explained by the use of molecular diagnostic methods used by et al., 2013). The lineage found in the sample GERPH: PC005 was Pineda-Catalan et al. (2013). These authors detected low parasitaemia closely related to the Hae. pacayae sequences identified as KF049495, by microscopy and a maximum of eight parasitic forms by blood smear and KF0449507 (group 2, with a mean distance within the group of (Pineda-Catalan et al., 2013). Low parasitemia was also detected in our 0.003 ± 0.001; Fig. 3C, and Supplementary Fig. S1). Genetic distances samples. However, estimating prevalence through microscopy can un- of 0.02 ± 0.006 were obtained between GERPH: PC005 and sequences derestimate the true rates of infection, mainly in cases of chronic (Jarvi named as Hae. pacayae s.l., group 1 (KF049506, KF049509, and et al., 2002) or mild infection. KF049513). As a comparison, the genetic distance within all available The prevalence reported here is low, even when compared to similar sequences identified as Hae. pacayae s.l. is an order of magnitude works that use microscopy as the only detection method such as in the greater than the ones estimated using the haplotypes for other well- case of Pe. dumerilianus, where a 50% prevalence was reported (Lainson known species like Hae. kopki, Hae. ptyodactylii, Hae. mesnili, Hae. and Naiff, 1998). In other organisms such as geckos, the reported pre- anatolicum and Hae. caucasica (see Fig. 3C, Supplementary Fig. S1). valence is comparable to this study's findings, being 6.7% in Lygo- Given that the genetic distance between GERPH: PC005 lineage and the dactylus capensis grotei (Telford, 2005). The parasitaemia observed in sequences named as Hae. pacayae s.l., group 1 (KF049506, KF049509, the current study is low compared to those reported for Omani geckos and KF049513) was 0.02 ± 0.006, the lineage GERPH: PC005 could be (0.3%, Maia et al., 2016). Despite this, our results are considerably considered as a new species; however, there is still not enough mole- lower when compared to the parasitaemias reported in Hae. lygodactyli cular data that can support this hypothesis. Nevertheless, a detailed in acute (56.4%) and chronic infection (20.2%) (Telford, 2005)orin description of this parasite is given. Elseya latisternum (between 1 and 50%) (Jakes et al., 2001). Low parasitaemia could prevent the uptake of the vector of the two sexual 3.3. Morphological description Haemocystidium (Simondia) sp. forms of gametocytes during feeding, thus affecting the transmission of the parasite. Although the tabanid fly Chrysops callidus (Diptera: Ta- Parasites do not induce hypertrophy of the red blood cells (mor- banidae) was shown as a vector for Hae. metchnikovi in the turtle phometrics have been provided in Table 2). The morphological de- Chrysemys picta in North America (DeGiusti et al., 1973), the vectors of scription is based on GERPH: PC005. these blood parasites in South America are unknown. Studies aimed at identifying the vector could help to decipher the host-parasites re- 3.3.1. Young gametocytes lationship between Haemocystidium and Podocnemis. The outline of gametocytes is not amoeboid (even); the shape varies Of the eight extant species of the Podocnemididae (Rhodin et al., from circular (33%), oval (30%) or ellipsoid (37%). The presence of a 2017), Podocnemis vogli has a relatively narrow distribution; never- large vacuole can be observed in the youngest gametocytes (Fig. 4A–D). theless, it is sympatric with Podocnemis expansa, Podocnemis unifilis and As the gametocyte develops, it acquires a doughnut shape (Fig. 4B–E), Peltocephalus dumerilianus in several regions of the Amazon and

304 L.P. González, et al. IJP: Parasites and Wildlife 10 (2019) 299–309

Table 2 Morphometrical features of Haemocystidium (Simondia) complex species described in South America and their hosts. Measurements of infected and non-infected erythrocytes are shown.

Characteristic Hae. (S) PC004 and PC006 Hae. (S) sp. PC005 Hae. (S.) pacayae Hae. (S.) peltocephali H. peltocephali (Lainson and Pineda-Catalan et al. Pineda-Catalan et al. Naiﬀ, 1998) (2013) (2013)

Host Podocnemis vogli P. expansa P. uniﬁlis Peltocephalus dumerilianu

This study

Non-infected erythrocytes n=15 n=15 ––– Length 22.5 ± 1.4 20.51 ± 2.92 ––– (18.9–24.6) (18.97–26.02) ––– Wide 14.1 ± 1.2 14.86 ± 1.33 ––– (11.4–16.0) (11.79–17.46) ––– Area 251.3 ± 28.1 258.86 ± 21.14 ––– (214.5–310.6) (219.63–292.91) ––– Nucleus Length 5.9 ± 0.7 5.66 ± 0.45 ––– (5.0–7.2) (5.05–6.12) ––– Wide 4.2 ± 0.5 4.03 ± 0.39 ––– (3.7–5.3) (3.71–4.52) ––– Area 20.8 ± 2.7 18.30 ± 4.05 ––– (15.9–26.4) (12.86–27.66) ––– Microgametocyte n=1 n=4 n=12 n=6 Length 10.44 8.44 ± 1.29 10.86 ± 2.30 10.11 ± 0.97 6.7–12.5 – (7.56–10.30) (5.91–14.13) (9.01–11.54) Wide 7.14 4.84 ± 0.44 6.87 ± 0.94 8.81 ± 1.04 6.0–9.0 – (4.46–5.47) (5.36–8.31) (7.01–9.91) – Area 67.54 41.09 ± 6.61 64.64 ± 19.77 74.73 ± 17.32 – – (36.24–50.29) (26.67–92.74) (62.16–98) – No Pigment granules > 10 > 10 Aggregated ––12–30 n=5 n=3 ––Average 21 0.42 ± 0.08 0.31 ± 0.11 ––n=17 (0.30–0.53) (0.16–0.54) –– Number of vacuoles 0.58 ± 1 0 Nucleus Length 5.42 –– – – ––––– Wide 0.65 –– – – ––––– Area 0.35 –– – – ––––– Macrogametocytes n =2 n=4 n=19 n=19 Length 9.45 ± 0.05 10.63 ± 1.23 11.03 ± 3.16 10.07 ± 1.77 7.4–12.6 (9.41–9.48) (9.38–11.55) (5.22–18.85) (6.15–14.07) Wide 6.12 ± 0.19 5.94 ± 0.95 7.68 ± 2.25 8.95 ± 1.41 6.2–11.1 (5.99–6.26) (5.26–7.23) (4.70–12.76) (5.69–11.11) Area 47.56 ± 2.88 52.01 ± 11.61 73.31 ± 32.29 76.84 ± 24.34 – (45.52–49.59) (40.47–65.89) (22.92–134.83) (28.11–130.92) Nº. Maximum of pigment > 10 > 10 Aggregated ––15–20 granules n=6 n=8 –– 0.28 ± 0.03 0.27 ± 0.05 –– (0.20–0.31) (0.21–0.38) –– Number of vacuoles 1.25 ± 0.5 (1–2) ––3 Nucleus n =2 n=3 ––– Length 5.78 ± 2.19 2.79 ± 0.76 ––– (4.23–7.34) (2.25–3.33) ––– Wide 0.67 ± 0.32 1.26 ± 0.16 ––– (0.44–0.89) (1.14–1.37) ––– Area 3.12 ± 0.03 3.46 ± 1.38 ––– (3.10–3.14) (2.18–5.27) ––– Infected erythrocytes n=3 n=8 ––– Length 23.40 ± 2.16 22.01 ± 2.12 ––– (20.43–26.38) (19.67–25.53) ––– Wide 14.07 ± 0.61 14.71 ± 1.41 ––– (13.32–14.74) (11.81–15.98) ––– Area 261.32 ± 21.00 262.88 ± 33.21 ––– (224.95–325) (219.18–325.06) ––– Nucleus Length 5.63 ± 0.23 5.61 ± 0.82 ––– (5.30–5.90) (4.53–6.74) ––– Wide 4.24 ± 0.29 4.04 ± 0.41 ––– (3.95–4.67) (3.63–4.60) ––– Area 19.41 ± 0.76 19.54 ± 2.24 ––– (18.51–20.33) (17.41–24.45) –––

305 L.P. González, et al. IJP: Parasites and Wildlife 10 (2019) 299–309

Orinoquia (Rueda-Almonacid et al., 2007), which could explain, at least unlike species of Haemoproteus parasitizing birds or Haemoproteus in part, the presence of Haemocystidium spp. in this family of turtles. It is mesnili (described in cobras). However, as shown in Fig. 3A, the three important to emphasize that the species belonging to the Podocnemi- species mentioned above are part of the same clade, suggesting that, didae share habitat with species of other families of semi-aquatic turtles despite the difference in merogony, these species seem to belong to the such as Kinosternon scorpioides (Kinosternidae) and also with tortoises proposed subgenus Haemocystidium. such as Chelonoidis carbonaria and Ch. denticulata (Testudinidae), which The results obtained in this study contrast with those published by were negative for microscopic infection by this parasite, however, the Boundenga et al. (2016, 2017), where the authors reported the presence sampling of these species was limited. Based on the data analyzed to of haemosporidian in K. erosa and Pel. castaneus in Gabon, Africa. In date, it would appear that Haemocystidium shows specificity for Po- these studies, Haemocystidium appears as a sister clade of the genera docnemididae species with the cis-Andean distribution. The absence of Leucocytozoon and Haemoproteus (Parahaemoproteus). Likewise, new detectable Haemocystidium infections in Podocnemis lewyana (n = 17) species of Haemoproteus have been described in birds, such as Haemo- and Trachemys venusta callirostris (n = 11), which are distributed along proteus catharti (from the Turkey vulture) and Haemoproteus sp. from the basins of the Magdalena and Sinú rivers in Colombia, and in trans- Mycteria americana (Wood Stork), which constitutes a separate clade Andean individuals of Chelonoidis carbonaria (n = 6) and Kinosternon from the other Haemoproteus spp. of birds. These two Haemoproteus scorpioides (n = 3), could be associated with the distribution of these species appear closely related to the genus Plasmodium in this study. turtle species and the absence of the possible vector. Lainson and Naiff This indicates the need to extend the sampling to new taxa of both birds (1998) reported the presence of Hae. geochelonis in a single individual and reptiles in order to improve the resolution of the phylogenetic re- Ch. denticulata. More testudinids from both sides of the Andean cor- lationships of haemosporidian parasites. dilleras should be screened to test this hypothesis that Haemocystidium It is important to highlight that the sequences identified in the in- shows specificity for Podocnemididae species with the cis-Andean dis- dividuals GERPH: PC004 and GERPH: PC006 are identical to Hae. pa- tribution. In birds, for example, the Leucocytozoon distribution in Co- cayae (KF049506) reported by Pineda-Catalan et al. (2013), extending lombia is restricted to the highlands (Matta et al., 2014; Lotta et al., the geographic and host range for this species. Haemocystidium (Si- 2016), probably also associated with the abundance and distribution of mondia) sp. (GERPH: PC005) is morphologically indistinguishable from the vectors. Haemocystidium peltocephali (KF049492) and Hae. pacayae (KF049506) The low prevalence of haemosporidian (1.55%) reported in this (Pineda-Catalan et al., 2013); however, the genetic distance of GERPH: study contrasts with the high prevalence of haemogregarines (84.53%) PC005 with these latter species, was 0.044 and 0.02, respectively found in the same group of samples analyzed. Yet, for the latter, the (Fig. 3B). Pineda-Catalan et al. (2013) reported several lineages for Hae. vector is also unknown; however, leeches as vectors have been reported pacayae, among them the lineages KF049495 and KF049507; however, before (Telford, 2009). Our results are consistent with other studies with these latter lineages, the genetic distance with GERPH: PC005 is conducted in testudines, where the most commonly reported parasite is only 0.006 (Fig. 3B). For that reason, although the lineages KF049495 Haemogregarina sp. (Rossow et al., 2013; Soares et al., 2014; Úngari and KF049507 were previously assigned as Hae. pacayae, our results et al., 2018). suggested that both of these lineages and GERPH: PC005 could be an- Concerning haemoproteids (Haemocystidium and previously named other Haemocystidium (Simondia) species; however, more molecular Haemoproteus), 16 species have been reported to date in reptiles markers are needed in order to support this hypothesis. (Supplementary Table 1). The genus Haemocystidium is characterized by There are only reports of Haemocystidium sp. for Pe. dumerilianus and the presence of malarial pigment (a trait shared with species of the Podocnemis spp. (distributed in the North of South America) among the genera Plasmodium and Haemoproteus) and the absence of peripheral three genera belonging to the family Podocnemididae. Characteristics blood erythrocytic meronts (a trait that it shares with the genus Hae- such as the presence/absence of vacuoles, nucleophile growth of the moproteus, but that set it apart from species of the genus Plasmodium). parasite, presence of a parasitophorous vacuole (PV), the distribution of Regardless of these differences, species of Haemocystidium in reptiles granules in the cytoplasm of macro- and microgametocytes and the have been described as Haemoproteus spp. in the past (Lainson and presence of amoeboid gametocytes, can be used as features that support Naiff, 1998; Jakes et al., 2001; Javanbakht et al., 2015). However, the the diagnosis in this genus (e.g., to gametocytes Hae. peltocephali in phylogenetic analysis of the cytb gene sequences previously reported peripheral blood). Nevertheless, species differentiation and identifica- (Pineda-Catalan et al., 2013; Maia et al., 2016) and those obtained in tion are difficult, as many characters overlap. Species described in other this study (Fig. 3), showed that Haemoproteus species infecting birds reptiles may be differentiated from those described in Podocnemidids and parasites of the genus Haemocystidium are two distinct mono- due to their size, where macro and microgametocytes surround the phyletic groups. However, limited sampling, that results in the lack of nucleus of the host cell in Testudines: H. metchnikovi (Simond, 1901), H. taxa from reptilian hosts, does not allow for the elucidation of whether testudinalis (Cook et al., 2010) synonyms Haemamoeba testudinis these are two monophyletic groups that support the subdivision of the (Laveran, 1905) and H. chelodinae (Mackerras, 1961; Jakes et al., 2001); genus Haemocystidium into two subgenera: Simondia infecting chelo- in snakes: H. mesnili (Telford, 2007) and in lizard: Hae. papernae, Hae. nians and Haemocystidium isolated from the Squamata as suggested by quettaensis (Telford, 1996), and H. lygodactyli (Telford, 2005). Pineda-Catalan et al. (2013) and Maia et al. (2016). Although we found The species concept in haemosporidian is still a matter of debate. that the turtle parasites could form a monophyletic group (Simondia), The concomitant use of morphological, morphometric, and genetic the host information of the Haemocystidium sp. (MH177855), included tools is ideal at present for addressing this issue (Martinsen et al., 2006; in this study's phylogenetic analyses, is not available, so it is not pos- Hellgren et al., 2007; Perkins et al., 2011; Pacheco et al., 2013; Outlaw sible to explore this issue with the available data. and Ricklefs, 2014). To date, the delimitation of species continues to be Nevertheless, our results suggest that species of Haemocystidium controversial, e.g. for the genus Haemoproteus (in birds), Hellgren et al. described in turtles appear to be part of four differentiated clades ac- (2007) suggested that cytb lineages with a genetic distance > 5% are cording to their geographical distribution: a clade formed by the species associated with morphologically differentiable species; however, the reported in Iran (Testudo sp.), another by the African species (Kinixys same authors report that the species H. minutus and H. pallidus (mor- erosa and Pelusios castaneus), and two clades with the species identified phologically differentiable) differ by 0.7%, while lineages associated in South America (Podocnemis sp.) (Fig. 3A, Supplementary Fig. S1). with morphospecies H. balmorali show a difference of 2.7%. Recently, Regarding the clade of species that infect lizards and snakes, Telford Galen et al. (2018) through an integrative approach, found cryptic (1996), analyzed tissues infected with Hae. kopki and Hae. ptyodactylii, species of Leucocytozoon with a minimum difference in a single base supporting the status of the genus Haemocystidium. This was based on pair in mDNA (approx. 0.2% divergence), and genetic distances for the the meronts of these species not forming pseudocytomers, which is nuclear markers between 1.07 and 5.19%.

306 L.P. González, et al. IJP: Parasites and Wildlife 10 (2019) 299–309

Table 3 Genetic distances reported in haemosporidian species, using cytb gene or mtDNA. Presence/absence of cryptic species is indicated by yes or no.

Authors Genetic Markers Size fragment Species 1 Species 2 Host Distance Cryptic species

Escalante et al. (1998) cytb 1035 bp Plasmodium gallinaceum Plasmodium elongatum Specie 1: Gallus gallus 0.034 No Specie 2: Passer domesticus Perkins (2000) cytb 540 bp Plasmodium azurophilum Plasmodium azurophilum Anolis gundlachi 0.031 Yes red blood cell while blood cell Palinauskas et al. cytb 479 bp Plasmodium Plasmodium circumﬂexum Species 1: 0.055 Yes (2015) homocircumﬂexum Lanius collurio Species 2: Acrocephalus scirpaceus Valkiūnas et al. (2010) cytb 706 bp Leucocytozoon mathisi Leucocytozoon buteonis Species 1: 0.109 Yes Accipiter nisus Species 2: Buteo jamaicensis Pacheco et al. (2013) Complete Species 1: Plasmodium falciparum Plasmodium reichenowi Species 1: 0.025 No mtDNA 5949 bp Homo sapiens Species 2: Species 2: 5966 bp Pan troglodytes Complete Species 1: Plasmodium vivax Plasmodium cynomolgi Species 1: 0.012 No mtDNA 5990 bp Homo sapiens Species 2: Species 2: 5991 bp M. radiata, Presbytis entrellus, Mantilla et al. (2013) Complete Species 1: Plasmodium lutzi Plasmodium relictum Species 1: 0.018 No mtDNA 5889 bp Turdus fuscater Species 2: Species 2: 6003 bp Zenaida macroura Walther et al. (2014) cytb Species 1: Plasmodium homopolare Plasmodium Species 1: 0.036 No 478 bp parahexamerium Melospiza melodia Species 2: Species 2: 468 bp Alethe diademata Muehlenbein et al. Complete Species 1: Plasmodium. knowlesi Plasmodium coatneyi Species 1: 0.032 No (2015) mtDNA 5957 bp Macaca fascicularis Species 2: Species 2: 5976 bp Macaca mulatta Maia et al. (2016) cytb Species 1: Hae. sp. (S7155) Hae. sp. (EU254531) Species 1: 0.074 No Species 2: Hemidactylus luqueorum 607 bp Species 2: Ergenia stokesii van As et al. (2016) cytb 500 pb Plasmodium. zonuriae Plasmodium intabazwe Species 1: 0.034 No Cordylus vittifer Species 2: Pseudocordylus melanotus Matta et al. (2018) cytb 1039 bp Plasmodium carmelinoi Plasmodium kentropyxi Species 1: 0.02 No Ameiva ameiva Species 2: Cnemidophorus cf. gramivagus

Descriptions of parasite species based on morphology, host, or life- the genus Plasmodium in lizards, species with a genetic distance of 0.03 history traits have been shown to be ambiguous and inconsistent from may be indicative of different species, despite their morphological si- the early days of molecular phylogenetics in Haemosporida (e.g., milarity (Perkins, 2000). Even in Plasmodium species, which are mor- Escalante et al., 1998). Nowadays, an integrative taxonomic approach phologically differentiable, there are reports of genetic distances, which using morphometry, nuclear and plastids molecular markers, as well as vary from 0.012 to 0.034 (Escalante et al., 1998; Pacheco et al., 2013). ecological features, are essential tools in the description of these Table 3 shows the published genetic differences for cryptic and non- parasite species; in birds, for example, parasite morphometrical char- cryptic species, showing high variability in their distances. Based on the acteristics have been shown to vary according to the host (e.g. Plas- above, we suggest that the genus Haemocystidium infecting Podocne- modium lutzi initially reported in Aramides cajaneus and later isolated in midids could be a complex of cryptic species. Turdus fuscater (Mantilla et al., 2013); both quite different in morpho- Considering the above, the genus Haemocystidium is perhaps one of metrical measurements). The analyses of molecular markers have al- the most representative examples of the importance of morphological lowed elucidation of a large number of cryptic species (Table 3), and molecular characterization, especially when considering the diffi- however, those studies that use only molecular tools, usually lose va- culties in the differentiation of species based solely on their morpho- luable information such as the detection of co-infections and abortive logical characters. Also, we invite to extend studies on the character- infections (false positives), as well as morphological characters ization of life cycles of haemoparasites in reptiles, and the generation of (Valkiūnas et al., 2008). new knowledge that contributes to the elucidation of the phylogenetic Currently, an association has been sought between the results of relationships of haemosporidian parasites and their vectors. morphological, morphometric and genetic information analyses; this is ū how Hellgren et al. (2007) and Valki nas et al. (2009) suggested that 5. Conclusions species belonging to the genus Haemoproteus with a genetic distance of ff 0.05 in the cytb gene correspond to di erent species. Nevertheless, The molecular lineages linked to morphological characteristics and species such as Haemoproteus minutus and Haemoproteus pallidus (mor- the phylogenetic analyses reported here support that Haemocystidium phologically distinguishable) have only a genetic distance of 0.01. For genus (Castellani, and Willey, 1904) is a monophyletic group separated

307 L.P. González, et al. IJP: Parasites and Wildlife 10 (2019) 299–309 from other haemosporidian groups. Our study extends the geographical mitochondrial genome. Proc. Natl. Acad. Sci. U.S.A. 95, 8124–8129. and host range for the genus in South America. Future studies are ne- Gaffney, E.S., Meylan, P.A., Wood, R.C., Simons, E., De Almeida Campos, D., 2011. ff Evolution of the side-necked turtles: the family Podocnemididae. Bull. Am. Mus. Nat. cessary to identify the e ects of the infection, its prevalence in the Hist. vertebrate host, and the vectors associated with its transmission. To Galen, S.C., Nunes, R., Sweet, P.R., Perkins, S.L., 2018. Integrating coalescent species date, the species of Haemocystidium, (at least in Podocnemidids) cannot delimitation with analysis of host specificity reveals extensive cryptic diversity debe differentiated using solely morphological characters. Therefore, ad- spite minimal mitochondrial divergence in the malaria parasite genus Leucocytozoon. BMC Evol. Biol. 18 (1), 128. ditional genetic and ecological information is needed to assess the di- Gouy, M., Guindon, S., Gascuel, O., 2010. SeaView version 4: a multiplatform graphical versity of Haemocystidium and to support species delimitation. user interface for sequence alignment and phylogenetic tree building. Mol. Biol. Evol. 27, 221–224. https://doi.org/10.1093/molbev/msp259. Hellgren, O., Križanauskiene, A., Valkiūnas, G., Bensch, S., 2007. Diversity and phylogeny Declaration of competing interest of mitochondrial cytochrome b lineages from six morphospecies of avian Haemoproteus (Haemosporida: Haemoproteidae). J. Parasitol. 93, 889–896. https:// The authors declare that they have no conflict of interest. doi.org/10.1645/GE-1051R1.1. Herrera, J.R., 2008. Estudio patológico retrospectivo de mortalidad en reptiles del zoológico Jaime Duque entre el año 1991 y el 2006. Universidad de la Salle, Acknowledgments Bogotá, D.C. Jakes, K.A., O'Donoghue, P., Munro, M., Adlard, R., 2001. Hemoprotozoa of freshwater turtles in Queensland. J WILDLIFE DIS 37, 12–19. https://doi.org/10.7589/0090- This study was supported by the Departamento Administrativo de 3558-37.1.12. Ciencias, Tecnología e Innovación COLCIENCIAS, Colombia Project Jarvi, S.I., Schultz, J.J., Atkinson, C.T., 2002. PCR diagnostics underestimate the pre- code 1101-776-57872. The authors wish to thank the members of the valence of avian malaria (Plasmodium relictum) in experimentally-infected passerines. J. Parasitol. 88, 153–158. https://doi.org/10.1645/0022-3395. GERPH group, the inter-institutional agreement between Unitrópico Javanbakht, H., Kvičerová, J., Dvořáková, N., Mikulíček, P., Sharifi, M., Kautman, M., and Parex Resources “Huella Galápaga", the University Foundation - Maršíková, A., Široký, P., 2015. Phylogeny, diversity, distribution, and host specifi- Unitrópico, and the support of the inhabitants of the Martínez-Parales city of Haemoproteus spp. (Apicomplexa: Haemosporida: Haemoproteidae) of pa- – family area. We thank Ariana Cristina Pacheco for the design of the laearctic tortoises. J. Eukaryot. Microbiol. 62, 670 678. https://doi.org/10.1111/jeu. 12227. silhouette. Kumar, S., Stecher, G., Tamura, K., 2016. MEGA7: molecular evolutionary genetics analysis version 7.0 for bigger datasets. Mol. Biol. Evol. 33, 1870–1874. https://doi. Appendix A. Supplementary data org/10.1093/molbev/msw054. Lainson, R., 2012. Atlas of Protozoan Parasites of the Amazonian Fauna of Brazil, vol. 1 Haemosporida of reptiles. Supplementary data to this article can be found online at https:// Lainson, R., Landau, I., Shaw, J.J., 1974. Observations on non-pigmented haemosporidia doi.org/10.1016/j.ijppaw.2019.10.003. of Brazilian lizards, including a new species of Saurocytozoon in Mabuya mabouya (Scincidae). Parasitology 69, 215–223. Lainson, R., Naiff, R.D., 1998. Haemoproteus (Apicomplexa: Haemoproteidae) of tortoises References and turtles. Proc. Biol. Sci. 265, 941–949. Lavern, A., 1905. Sur une hëmamibe nouvelle de Testudo pardalis. C. R. Soc. Sëanc. Biol. 59, 176–178. š Adl, S.M., Bass, D., Lane, C.E., Luke , J., Schoch, C.L., Smirnov, A., Agatha, S., Berney, C., Levine, N.D., 1988. The Protozoan Phylum Apicomplex. CRC Press, INC, Boca Raton, Č č Brown, M.W., Burki, F., Cárdenas, P., epi ka, I., Chistyakova, L., del Campo, J., Florida, pp. 118–133 1988. Dunthorn, M., Edvardsen, B., Eglit, Y., Guillou, L., Hampl, V., Heiss, A.A., Levine, N.D., 1970. Taxonomy of the sporozoa. J. Parasitol. 56, 208–209. Hoppenrath, M., James, T.Y., Karpov, S., Kim, E., Kolisko, M., Kudryavtsev, A., Lahr, Lotta, I.A., Pacheco, M.A., Escalante, A.A., González, A.D., Mantilla, J.S., Moncada, L.I., D.J.G., Lara, E., Le Gall, L., Lynn, D.H., Mann, D.G., Massana i Molera, R., Mitchell, Adler, P.H., Matta, N.E., 2016. Leucocytozoon diversity and possible vectors in the E.A.D., Morrow, C., Park, J.S., Pawlowski, J.W., Powell, M.J., Richter, D.J., Rueckert, neotropical highlands of Colombia. Protist 167, 185–204. S., Shadwick, L., Shimano, S., Spiegel, F.W., Torruella i Cortes, G., Youssef, N., Mackerras, M., 1961. The haematozoa of Australian reptiles. Aust. J. Zool. 9, 61. https:// fi Zlatogursky, V., Zhang, Q., 2019. Revisions to the classi cation, nomenclature, and doi.org/10.1071/ZO9610061. diversity of eukaryotes. J. Eukaryot. Microbiol. https://doi.org/10.1111/jeu.12691. Maia, J.P., Harris, D.J., Carranza, S., 2016. Reconstruction of the evolutionary history of š Adl, S.M., Simpson, A.G.B., Lane, C.E., Luke , J., Bass, D., Bowser, S.S., Brown, M.W., Haemosporida (Apicomplexa) based on the cyt b gene with characterization of Burki, F., Dunthorn, M., Hampl, V., Heiss, A., Hoppenrath, M., Lara, E., le Gall, L., Haemocystidium in geckos (Squamata: gekkota) from Oman. Parasitol. Int. 65, 5–11. Lynn, D.H., McManus, H., Mitchell, E.A.D., Mozley-Stanridge, S.E., Parfrey, L.W., Mantilla, J.S., Matta, N.E., Pacheco, M.A., Escalante, A.A., González, A.D., Moncada, L.I., Pawlowski, J., Rueckert, S., Shadwick, L., Schoch, C.L., Smirnov, A., Spiegel, F.W., 2013. Identification of Plasmodium (Haemamoeba) lutzi (Lucena, 1939) from Turdus fi – 2012. The revised classi cation of eukaryotes. J. Eukaryot. Microbiol. 59, 429 514. fuscater (great thrush) in Colombia. J. Parasitol. 99, 662–668. https://doi.org/10.1111/j.1550-7408.2012.00644.x. Martinsen, E.S., Paperna, I., Schall, J.J., 2006. Morphological versus molecular identifi- ff Altschul, S.F., Madden, T.L., Schä er, A.A., Zhang, J., Zhang, Z., Miller, W., Lipman, D.J., cation of avian Haemosporidia: an exploration of three species concepts. Parasitology 1997. Gapped BLAST and PSI-BLAST: a new generation of protein database search 133, 279. https://doi.org/10.1017/S0031182006000424. – programs. Nucleic Acids Res. 25, 3389 3402. Matta, N.E., González, L.P., Pacheco, M.A., Escalante, A.A., Moreno, A.M., González, A.D., Bensch, S., Stjernman, M., Hasselquist, D., Ostman, O., Hansson, B., Westerdahl, H., Calderón-Espinosa, M.L., 2018. Plasmodium parasites in reptiles from the Colombia fi Pinheiro, R., 2000. Host speci city in avian blood parasites: a estudy of Plasmodium Orinoco-Amazon basin: a re-description of Plasmodium kentropyxi Lainson R, Landau fi and Haemoproteus mitochondrial DNA ampli eld from birds. Proc Biol Sci 267, I, Paperna I, 2001 and Plasmodium carmelinoi Lainson R, Franco CM, da Matta R, – 1583 1589. 2010. Parasitol. Res. 117, 1357–1370. https://doi.org/10.1007/s00436-018-5815-9. Benson, D.A., Karsch-Mizrachi, I., Clark, K., Lipman, D.J., Ostell, J., Sayers, E.W., 2012. Matta, N.E., Pacheco, M.A., Escalante, A.A., Valkiūnas, G., Ayerbe-Quiñones, F., Acevedo- – GenBank. Nucleic Acids Res. 40, D48 D53. https://doi.org/10.1093/nar/gkr1202. Cendales, L.D., 2014. Description and molecular characterization of Haemoproteus Boundenga, L., Makanga, B., Ollomo, B., Gilabert, A., Rougeron, V., Mve-Ondo, B., macrovacuolatus n. sp.(Haemosporida, Haemoproteidae), a morphologically unique Arnathau, C., Durand, P., Moukodoum, N.D., Okouga, A.-P., Delicat-Loembet, L., blood parasite of black-bellied whistling duck (Dendrocygna autumnalis) from South Yacka-Mouele, L., Rahola, N., Leroy, E., Ba, C.T., Renaud, F., Prugnolle, F., Paupy, C., America. Parasitol. Res. 113, 2991–3000. 2016. Haemosporidian parasites of antelopes and other vertebrates from Gabon, Morrison, D.A., 2009. Evolution of the Apicomplexa: where are we now? Trends Parasitol. Central Africa. PLoS One 11, e0148958. https://doi.org/10.1371/journal.pone. 25, 375–382. https://doi.org/10.1016/j.pt.2009.05.010. 0148958. Muehlenbein, M.P., Pacheco, M.A., Taylor, J.E., Prall, S.P., Ambu, L., Nathan, S., Alsisto, Boundenga, L., Perkins, S.L., Ollomo, B., Rougeron, V., Leroy, E.M., Renaud, F., Prugnolle, S., Ramirez, D., Escalante, A.A., 2015. Accelerated diversification of nonhuman pri- F., 2017. Haemosporidian parasites of reptiles and birds from Gabon, Central Africa. mate malarias in southeast Asia: adaptive radiation or geographic speciation? Mol. – J. Parasitol. 103, 330 337. https://doi.org/10.1645/16-118. Biol. Evol. 32, 422–439. https://doi.org/10.1093/molbev/msu310. Castaño-Mora, O.V., 2002. Libro rojo de reptiles de Colombia. La serie Libros rojos de Outlaw, D.C., Ricklefs, R.E., 2014. Species limits in avian malaria parasites especies amenazadas. Universidad Nacional de Colombia. (Haemosporida): how to move forward in the molecular era. Parasitology 141, Castellani, A., Willey, A., 1904. Observations on the haematozoa of vertebrates in Ceylon. 1223–1232. https://doi.org/10.1017/S0031182014000560. – Spolia Zeylonic 2, 78 92. Pacheco, M., Cranfield, M., Cameron, K., Escalante, A.A., 2013. Malarial parasite diversity Cook, C.A., Smit, N.J., Davies, A.J., 2010. Hemoproteids (Apicomplexa: Haemoproteidae) in chimpanzees: the value of comparative approaches to ascertain the evolution of – from South African tortoises (Cryptodira: Testudinidae). J. Parasitol. 96, 1168 1172. Plasmodium falciparum antigens. Malar. J. 12, 328. https://doi.org/10.1186/1475- https://doi.org/10.1645/GE-2527.1. 2875-12-328. DeGiusti, D.L., Sterling, C.R., Dobrzechowski, D., 1973. Transmission of the Chelonian Pacheco, M.A., Cepeda, A.S., Bernotienė, R., Lotta, I.A., Matta, N.E., Valkiūnas, G., fl haemoproteid Haemoproteus metchnikovi by a tabanid y Chrysops callidus. Nature Escalante, A.A., 2018a. Primers targeting mitochondrial genes of avian haemospor- – 242, 50 51. https://doi.org/10.1038/242050a0. idians: PCR detection and differential DNA amplification of parasites belonging to Escalante, A.A., Freeland, D.E., Collins, W.E., Lal, A.A., 1998. The evolution of primate different genera. Int. J. Parasitol. 48, 657–670. https://doi.org/10.1016/j.ijpara. malaria parasites based on the gene encoding cytochrome b from the linear 2018.02.003.

308 L.P. González, et al. IJP: Parasites and Wildlife 10 (2019) 299–309

Pacheco, M.A., Matta, N.E., Valkiūnas, G., Parker, P.G., Mello, B., Stanley, C.E., Lentino, Staats, C.M., Schall, JosJ., 1996. Distribution and abundance of two malarial parasites of M., Garcia-Amado, M.A., Cranfield, M., Kosakovsky Pond, S.L., Escalante, A.A., the endemic Anolis lizard of saba island, Netherlands antilles. J. Parasitol. 82, 2018b. Mode and rate of evolution of haemosporidian mitochondrial genomes: 409–413. https://doi.org/10.2307/3284077. timing the radiation of avian parasites. Mol. Biol. Evol. 35, 383–403. https://doi.org/ Tamura, K., Nei, M., 1993. Estimation of the number of nucleotide substitutions in the 10.1093/molbev/msx285. control region of mitochondrial DNA in humans and chimpanzees. Mol. Biol. Evol. Pachón, D.A., 2009. Aislamiento, identificación y serotipificación de enterobacterias del https://doi.org/10.1093/oxfordjournals.molbev.a040023. género Salmonella en una población Crocodylus intermedius y testudinos mantenidos Telford, S.R., 2005. A Haemocystidium species from the east african gecko Lygodactylus en cautiverio en la estación Biológica Roberto Franco E.B.T.R.B de la Facultad de capensis grotei. J. Parasitol. 91, 135–138. https://doi.org/10.1645/GE-3395. Ciencias- Universidad Nacional de Colombia. Pontificia Universidad Javeriana, Telford, S.R.J., 2007. Redescription of Haemoproteus mesnili (Apicomplexa: Plasmodiidae) Bogotá, D.C. and its meronts, with description of a second haemosporidian parasite of African Palinauskas, V., Žiegytė, R., Ilgūnas, M., Iezhova, T.A., Bernotienė, R., Bolshakov, C., Cobras. J. Parasitol. 93, 673–679. https://doi.org/10.1645/GE-3582.1. Valkiūnas, G., 2015. Description of the first cryptic avian malaria parasite, Telford Jr., S.R., 2009. Hemoparasites of the Reptilia: Color Atlas and Text. Taylor & Plasmodium homocircumflexum n. sp., with experimental data on its virulence and Francis, Boca Raton. development in avian hosts and mosquitoes. Int. J. Parasitol. 45, 51–62. https://doi. Telford Jr., S.R., 1996. Two new species of Haemocystidium Castellani & Willey org/10.1016/j.ijpara.2014.08.012. (Apicomplexa: Plasmodiidae) from Pakistani lizards, and the support their meronts Perkins, 2000. Species concepts and malaria parasites: detecting a cryptic species of provide for the validity of the genus. Syst. Parasitol. 34, 197–214. https://doi.org/10. Plasmodium. P ROY SOC B-BIOL SCI267, 2345–2350. https://doi.org/10.1098/rspb. 1007/BF00009387. 2000.1290. Úngari, L.P., Santos, A.L.Q., O'Dwyer, L.H., da Silva, M.R.L., de Melo Fava, N.N., Paiva, Perkins, S.L., Martinsen, E.S., Falk, B.G., 2011. Do molecules matter more than mor- G.C.M., de Melo Costa Pinto, R., Cury, M.C., 2018. Haemogregarina podocnemis sp. phology? Promises and pitfalls in parasites. Parasitology 138, 1664–1674. https:// nov.: description of a new species of Haemogregarina Danilewsky 1885 (Adeleina: doi.org/10.1017/S0031182011000679. haemogregarinaidae) in free-living and captive yellow-spotted river turtles Perkins, S.L., 2014. Malaria's many mates: past, present, and future of the systematics of Podocnemis unifilis (Testudines: Podocnemididae) from Brazil. Parasitol. Res. 117, the order Haemosporida. J. Parasitol. 100 (1), 11–26. https://doi.org/10.1645/13- 1535–1548. https://doi.org/10.1007/s00436-018-5817-7. 362.1. Valkiūnas, G., Iezhova, T.A., 2018. Keys to the avian malaria parasites. Malar. J. 17, 212. Pineda-Catalan, O., Perkins, S.L., Peirce, M.A., Engstrand, R., Garcia-Davila, C., Pinedo- https://doi.org/10.1186/s12936-018-2359-5. Vasquez, M., Aguirre, A.A., 2013. Revision of hemoproteid genera and description Valkiūnas, G., Iezhova, T.A., Križanauskienė, A., Palinauskas, V., Sehgal, R.N.M., Bensch, and redescription of two species of Chelonian hemoproteid parasites. J. Parasitol. 99, S., 2008. A comparative analysis of microscopy and PCR-based detection methods for 1089–1098. https://doi.org/10.1645/13-296.1. blood parasites. J. Parasitol. 94, 1395–1401. https://doi.org/10.1645/GE-1570.1. Rhodin, A.G.J., Iverson, J.B., Bour, R., Fritz, U., Georges, A., Shaffer, H.B., van Dijk, P.P., Valkiūnas, G., Iezhova, T.A., Loiseau, C., Sehgal, R.N.M., 2009. Nested cytochrome B 2017. Turtles of the world: annotated checklist and atlas of taxonomy, synonymy, polymerase chain reaction diagnostics detect sporozoites of hemosporidian parasites distribution, and conservation status. In: Conservation Biology of Freshwater Turtles in peripheral blood of naturally infected birds. J. Parasitol. 95, 1512–1515. https:// and Tortoises: a Compilation Project of the IUCN/SSC Tortoise and Freshwater Turtle doi.org/10.1645/GE-2105.1. Specialist Group. Chelonian Research Monograph, eighth ed. https://doi.org/10. Valkiūnas, G., Sehgal, R.N.M., Iezhova, T.A., Hull, A.C., 2010. Identification of 3854/crm.7.checklist.atlas.v8.2017. Leucocytozoon toddi group (Haemosporida: Leucocytozoidae), with remarks on the Ronquist, F., Huelsenbeck, J.P., 2003. MrBayes 3: Bayesian phylogenetic inference under species taxonomy of leucocytozoids. J. Parasitol. 96, 170–177. https://doi.org/10. mixed models. Bioinformatics 19, 1572–1574. https://doi.org/10.1093/ 1645/GE-2109.1. bioinformatics/btg180. van As, J., Cook, C.A., Netherlands, E.C., Smit, N.J., 2016. A new lizard malaria parasite Rossow, J.A., Hernandez, S.M., Sumner, S.M., Altman, B.R., Crider, C.G., Gammage, M.B., Plasmodium intabazwe n. sp. (Apicomplexa: haemospororida: Plasmodiidae) in the Segal, K.M., Yabsley, M.J., 2013. Haemogregarine infections of three species of aquatic Afromontane Pseudocordylus melanotus (Sauria: Cordylidae) with a review of African freshwater turtles from two sites in Costa Rica. Int J Parasitol Parasites Wildl 2, saurian malaria parasites. Parasites Vectors 9. https://doi.org/10.1186/s13071-016- 131–135. https://doi.org/10.1016/j.ijppaw.2013.02.003. 1702-3. Rueda-Almonacid, Carr, J.L., Mittermeier, R.A., Rodríguez-Mahecha, J.V., Mast, R.B., Vargas-Ramirez, M., Castaño-Mora, O., Fritz, U., 2008. Molecular phylogeny and diver- Vogt, R.C., Mittermeier, C.G., 2007. Las tortugas y los cocodrilianos de los países gence times of ancient South American and Malagasy river turtles (Testudines: andinos del trópico. Serie de guías tropicales de campo 6, 412–423. pleurodira: Podocnemididae). Org. Divers. Evol. 8, 388–398. https://doi.org/10. Sambrook, J., Fritsch, E.F., Maniatis, T., 1989. Molecular Cloning. Cold spring harbor 1016/j.ode.2008.10.001. laboratory press, New York. Walther, E.L., Valkiūnas, G., González, A.D., Matta, N.E., Ricklefs, R.E., Cornel, A., Schneider, C.A., Rasband, W.S., Eliceiri, K.W., 2012. NIH Image to ImageJ: 25 years of Sehgal, R.N.M., 2014. Description, molecular characterization, and patterns of dis- image analysis. Nat. Methods 9, 671–675. https://doi.org/10.1038/nmeth.2089. tribution of a widespread New World avian malaria parasite (Haemosporida: Simond, P.L., 1901. Contribution á l'ëtude des hëmatozoaires endoglobulaires des reptils. Plasmodiidae). Plasmodium (Novyella) homopolare sp. nov. Parasitol. Res 113, Ann. Inst. Pasteur. 15, 319–351. 3319–3332. https://doi.org/10.1007/s00436-014-3995-5. Soares, P., de Brito, E.S., Paiva, F., Pavan, D., Viana, L.A., 2014. Haemogregarina spp. in a WHO, 2018. Malaria [WWW Document]. world health organization. accessed 3.26.19. wild population from podocnemis unifilis troschel, 1848 in the Brazilian amazonia. https://www.who.int/es/news-room/fact-sheets/detail/malaria. Parasitol. Res. 113, 4499–4503.

309 Supplementary Table S1. List of species of Haemoproteus/Haemocystidium reported to the date infecting reptiles worldwide.

Author Country Species Host Chitra indica Chrysemys pictac Pseudemys elegansb Simond, 1901 India; North America H. metchnikovi Graptemys geographicac Emys blandingic Trionyx spiniferusc Stigmochelys pardalis Bouet, 1909 West Africa H. roumei (Cinixys belliana) Chelodina longicollis Australia H. chelodinae Johnston, and Cleland, Chelodina oblonga 1909 Australia Haemoproteus sp. Emydura krefftii Stauotypus triporcatus Chrysemys picta Kinixys homeana (Cinixys homeana) Belize; North America; Plimmer, 1912 Haemocystidium sp. Kinixys erosa. West Africa (Cinixys erosa) Kinixys belliana (Cinixys belliana)

Pittaluga, 1912 New Guinea H. cajali Clemmys Africana Chelodina longicollis C. oblonga Emydura macquarii Hae. chelodinae E. latisternum E. kreffti Mackerras, 1961 Australia Elseya dentata Phyllurus platurus Heteronota binoei Hae simondi Oedura tryoni Gehyra variegata australis Krasilnikov, 1965 Georgia, Russia H. caucasica Testudo graeca Misra and Choudhury, India H. trionyxii Trionyx gangeticus 1977 Pakistan Hae. Papernain Agama nuptafusca Telford, 1996 Pakistan Hae. quettaensis Agama nupta nupta

H. geochelonis Geochelone denticulate Lainson and Naiff, Amazon Brazil 1998 H. peltocephali Peltocephalus dumerilianus Elseya latisternum Jakes et al., 2001 Australia H. chelodinae E. signata

Telford, 2005 Tanzania Hae. lygodactyli Lygodactylus capensis grotei H. degiusti Telford, 2009 North America Chrysemys picta marginata (syn. H. metchnikovi) H. testudinalis Kinixys lobatsiana Cook et al., 2010 South Africa (syn. of Haemamoeba Stigmochelys pardalis testudinis Laveran, 1905)

South Africa H. natalensis Kinixys natalensis Orkun and Güven, Turkey H. anatolicum Testuda graeca 2013 Pacaya Samiria Hae. (Simondia) Podocnemis unifilis National Reserve, pacayae Podocnemis expansa Loreto, Peru Pineda-Catalan et al., Hae. (Simondia) 2013 Rio Negro, near Podocnemis unifilis peltocephali Barcelos, Amazonas Podocnemis expansa Lainson and Naiff State, Brazil Peltocephalus dumerilianus 1998 Chelonoidis carbonaria Martinele et al., 2016 Brazil Haemoproteus sp. Chelonoides denticulata Afghanistan H. anatolicum Testudo graeca Georgia Javanbakht et al., 2015 Iran Testudo horsfieldii H. caucasica Turkey Testudo graeca Boundenga et al., 2016 Gabon, Africa Hae. sp. Kinixys erosa Pelusios castaneus Boundenga et al., 2017 Gabon, Africa Hae. sp. Kinixys erosa

Supplementary Table S2. Primers used in PCR assays to detect haemosporidian parasites in sampled turtles.

Nested- Identified Primer sequence Reference PCR/Primer code parasite OUTER AE974 5' - TGTAATGCCTAGAMGWATWCC -3' Pacheco et al., Haemosporidian AE299 5' - GTCAAWCAAACATGAATATAGAC - 3' 2018a INNER AE983 5' - TGGATHTGTGGWGGATATYTWG - 3' Pacheco et al., Plasmodium sp. AE985 5' - AACGACCATATAWAATGWADATATC - 3' 2018a

References

Bouet, J., 1909. Sur deux he ̈mocytozaires pigmente ̈s des reptiles. C.R. Se ̈anc. Soc. Biol. 66, 43–

45.

Johnston, T.H., Cleland, J.B., 1909. On a new melanin- producing haematozoon from an Australian

tortoise. J. Proc. R. Soc. NSW 43, 97–103.

Krasilnikov, E.N., 1965. Parazity krovy cherepakh yugo- vostochnoi Gruzii (Blood parasites of

turtles of southeast Georgia). Zool. Zh. 44, 1454–1459.

Martinele, I., Tostes, R., Castro, R., D’Agosto, M., 2016. Prevalence of Haemoproteus spp.

(Apicomplexa: Haemoproteidae) in tortoises in Brazil and its molecular phylogeny. Parasitol.

Res. 115, 249–254. https://doi.org/10.1007/s00436-015-4741-3 Misra, K.K., Choudhury, A., 1977. Haemoproteus trionyxii n.sp. from a river turtle, Trionyx

gangeticus Cuvier. Arch. Protistenkd. 119, 395–400.

Orkun, Ö., Güven, E., 2013. A New Species of Haemoproteus from a Tortoise (Testudo graeca ) in

Turkey, with Remarks on Molecular Phylogenetic and Morphological Analysis. J. Parasitol. 99,

112–117. https://doi.org/10.1645/GE-3100.1

Pittaluga, J., 1912. Ein neuer Blutparasit de afrikanischer Schildröte, Clemmys africana ,

Haemoproteus cajale n.sp. Centralbl. f. Bakt. Parasitkde 63, 241–243.

Plimmer, H.G., 1912. On the blood parasites found in animals in the Zoological Gardens during the

four years 1908-1911. Proc. Zool. Soc. Lond. 27, 406–419.

Supplementary Figure S1. A Bayesian phylogenetic hypothesis of reptile haemosporidian parasites based on 103 partial sequences of cytb gene sequences corresponding to a small fragment of cytb (233 bp excluding gaps).

IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

3. Capítulo 3. The puzzle of Haemogregarine infection in aquatic South American turtles. En Colombia se han reportado 28 especies de tortugas terrestres. La información relacionada con la presencia de hemoparásitos en este grupo es limitada, ya que se desconoce la diversidad real de estos organismos. Aquí, documentamos la presencia de especies de Haemogregrina, en los podocnemididos, Podocnemis vogli y P. unifilis en dos localidades de la Orinoquia colombiana. Los individuos analizados fueron capturados utilizando trampas de embudo, redes de pesca y a mano. Las muestras obtenidas no superaron el 1% del peso de los individuos. La presencia de hemoparásitos se evaluó mediante microscopía para determinar su morfología y mediante un protocolo de PCR dirigido al 18S rRNA. Se encontró una prevalencia del 76% de infectados con la identificación de tres morfos morfológicamente diferenciables. Sin embargo, las distancias genéticas obtenidas entre las diferentes secuencias, nos llevaron a creer que dos de estos morfotipos correspondían a diferentes etapas de maduración. Teniendo en cuenta las limitaciones del 18S rRNA, la variabilidad de las secuencias y un tercer morfotipo que no fue posible amplificar, sugerimos una búsqueda de nuevos marcadores que puedan mejorar la resolución de las relaciones filogenéticas de los parásitos del suborden Adeleorina. El presente estudio arroja luz sobre la distribución geográfica y de distribución de hospedantes para el género Haemogregarina, contribuyendo al conocimiento de la diversidad de hemoparásitos en la herpetofauna colombiana.

Palabras claves: Hemoparásito, reptil, Simondia, Quelonios, Colombia.

Este artículo abarca los tres objetivos del proyecto de maestría, donde se analizan los resultados obtenidos para Haemogregarinas a nivel morfológico y molecular, con el fin de caracterizar los morfotipos y linajes identificados. Mi contribución en estee manuscrito radicó en la revisión de los extendidos sanguíneos de los individuos incluidos en el estudio, la toma de fotografías y medidas, el análisis de las secuencias obtenidas, el análisis detallado de la morfología de cada uno de los morfotipos, determinar las parasitemias y la realización de los análisis filogenéticos. Ademas, de contribuir con la escritura del artículo.

Este manuscrito muestra los resultados parciales a la fecha, dados los resultados obtenidos con el marcador molecular 18S rRNA se consideró pertinente complementar dicha información con mayor profundidad. Se planea por tanto, que algunas de las sugerencias de este manuscrito sean abordadas y complementadas en la tesis de maestria del estudiante German Alfredo Gutierrez Liberato, de la maestria en infecciones y salud en el trópico. IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

The puzzle of Haemogregarine infection in aquatic South American turtles.

3.1 Abstract In Colombia 28 species of land tortoises have been reported. Information relating to the presence of haemoparasites in this group is limited since the actual diversity of these organisms is unknown. Here, we document the presence of Haemogregrina species, in the podocnemidids, Podocnemis vogli and P. unifilis in two localities of the Colombian

Orinoquia. The analyzed individuals were captured by using funnel traps, fishing nets and by hand. The samples obtained did not exceed 1% of the weight of the individuals. The presence of haemoparasites was evaluated by microscopy for their morphology and by a

PCR protocol directed to 18s RNA. A prevalence of 76% infected was found with the identification of three morphologically differentiable morphs. However, the genetic distances obtained among the different sequences, led us to believe that two of these morphotypes corresponded to different stages of maturation. Taking into account the limitations of the 18s RNA, the variability of sequences, and a third morphotype that was not possible to amplify, we suggest a search for new markers that could improve the resolution of phylogenetic relationships of the parasites of the suborder Adeleorina. The present study, shed lights on the geographic and host range distribution for the

Haemogregarina genus, contributing to the knowledge of the diversity of haemoparasites in the Colombian herpetofauna. 99 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

3.2 Introduction Turtles are one of the most threatened groups of vertebrate (Hoffmann et al. 2010). In

Colombia, there are nine families of turtles, 28 species are continental (aquatic and terrestrial), and six are marine species (Forero-Medina et al. 2014). Fifteen of these species are classified in different categories of threat in the IUCN categories (Morales-Betancourt et al. 2015). The main factors associated with the decline of turtle populations are illegal trafficking of species and the consumption of their meat and eggs (Ortiz-Moreno and

Rodríguez-Pulido 2017). Recently, there has been increasing interest in characterizing infections caused by parasites in wild herpetofauna (Picelli et al. 2015; de Oliveira et al.

2018) as well as those confined to zoos and rehabilitation centres (Zamudio and Ramírez

2007).

Blood parasites affect the fitness of their vertebrate hosts. Deleterious effects on the reproductive success of birds are documented (Merino et al. 2000), as well as weight loss

(Valkiūnas et al. 2006) and susceptibility to predation (García-Longoria et al. 2015). In reptiles, it has been reported that blood parasites can decrease fat storage (Schall 1983) and alter haematological parameters, such as a decrease in haemoglobin or an increase in immature red blood cells (Oppliger et al. 1996). However, in reptiles the haemoparasite- host association is apparently benign (Brown et al. 2006).

The family Podocnemididae (Rueda-Almonacid et al. 2007) originated in the Cretaceous, which at the time contained about 20 genera and 30 species. Currently, only three of these genera (eight species), are extant in South America (Podocnemis and Peltocephalus) and

Madagascar (Erymnochelys), the rest are extinct (Gaffney et al. 2011). For the genus

Peltocephalus the only species reported is P. dumerilianus. Podocnemis contains six species P. erythrocephala, P. expansa, P. lewyana, P. sextuberculata, P. unifilis, and P. Capítulo 3 100 vogli (Vargas-Ramirez et al. 2008) distributed in Colombia. Of these, P. expansa and P. vogli are the most basal species, having diverged in the Late Eocene and late

Oligocene.Podocnemis vogli (savanna side-necked turtle) is distributed in the Colombian-

Venezuelan Orinoquia, while Podocnemis unifilis (Yellow-spotted River Turtle), is distributed in the Amazon as well as the Orinoquia

Among the blood parasites most commonly reported in turtles are the Haemogregarina species (Davis and Sterrett 2011; Rossow et al. 2013; Soares et al. 2014; DvořáKová et al.

2015). These parasites belong to the Apicomplexa phylum, which groups obligate intracellular protozoa that infect birds, reptiles and mammals, with around 5000 species recognized (Morrison 2009). The order Eucoccidiorida contains the suborder Adeleorina to which belong the families Hepatozoidae (one genus Hepatozoon), Karyolysidae (Hemolivia and Karyolysus), Dactylosomatidae (Babesiosoma and Dactylosoma), Haemogregarinidae

(Haemogregarina, Desseria and Cyrilia), Adeleidae (7 genera), Klosiellidae and

Legerellidae (the latter three families infect marine invertebrates) (Adl et al. 2019). In reptiles, the presence of the genera Hepatozoon (infecting snakes), Karyolysus (infecting lizards), Hemolivia (infecting turtles and amphibians) and Haemogregarina (infecting turtles) is known (Telford 2009; Maia 2015).

The identification of organisms placed in the Adeleorina suborder is traditionally based on the morphological characteristics of the parasite; however, the absence of diagnostic characters, such as, sexual dimorphism and hemozoin granules makes the description of species problematics. The most commonly used marker for the identification of the haemoparasites belonging to the suborder Adeleorina is the 18S rRNA gene. The RNAr gene is a multigene family that in eukaryotes has multiple copies. The 18S rRNA gene has regions with different rates of evolution from highly conservative to highly variable (Morand et al. 2015). In contrast with other eukaryotes where the RNA units are clustered in tandem, 101 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA. in some Apicomplexa as Plasmodium, Theileria and Babesia are not grouped in tandem, instead they are disperse throughout the genome (Dalrymple 1990; Kibe et al. 1994;

McCutchan et al. 1995). The small subunit of rRNA (18S rRNA) specifically shows a slow rate of evolution and is highly conserved (Morand et al. 2015). Recently, new species of haemoparasites have been described based on analyses of phylogenies using fragments of the 18S rRNA gene such as Karyolysus paradoxa (Cook et al. 2016) and Hepatozoon ixoxo (Netherlands et al. 2014).

In general, the life cycle of the suborder Adeleorina involves a vertebrate host where merogony and gamogony occur, as well as in an invertebrate host where sporogony develops. Leeches are the vectors associated with the transmission of the genus

Haemogregarina (Siddall and Desser 1991; Telford 2009). The place where merogony occurs can vary, for example, in the genus Hepatozoon, this process has been reported to occur in the liver the parenchymal and endothelial cells, while in the genus Haemogregarina there is first merogony in pre-erythrocytic cells, liver, lungs and spleen and second merogony in the erythrocytes (Telford 2009).

The Haemogregarina genus is reported for countries in Europe, Asia, America, Australia and Africa (Laird 1950; Telford 2009; Soares et al. 2014; DvořáKová et al. 2015; Picelli et al. 2015). In turtles, genera such as Hemolivia (Široký et al. 2005; Paperna2006; Siroky et al. 2007), Haemogregarina (Siddall and Desser 1992; Davis and Sterrett 2011; Rossow et al. 2013; Úngari et al. 2018) and Haemoproteids (Lainson and Naiff 1998; Cook et al. 2010;

Orkun and Güven 2013) are known. In Colombia, studies have been conducted relating to various organisms that infect turtles (Zamudio and Ramírez 2007; Gutierrez 2016)

Recently, for example, Haemocystidium sp., was detected in the Colombian Orinoquia in

Podocnemis vogli (savanna side-necked turtle) (González et al., 2019) Capítulo 3 102

Using morphological and molecular information, this study addresses the prevalence and diversity of parasites of the genus Haemogregarina in the aquatic turtles the savanna side- necked turtle and the yellow-spotted river turtle.

3.3 Materials and methods

3.3.1 Sampling A total of 190 free-living turtles were sampled, representing nine species and six families

(Table III 1). Of these, 147 individuals belonged to Podocnemis vogli were collected in two rural locations in the departments of Casanare, Hato la Esperanza (April and October 2017:

May 2018) (n = 134/147) and Vichada-Colombia: Finca Las Flores (April 2017) (n = 13/147), respectively (Fig. III 1). One specimen of Podocnemis unifilis was collected in the department of Arauca (Cravo norte). The sampling sites are between 200 - 250 m.a.s.l with an average annual temperature between 26 - 28 ° C. The sampling period corresponded to the dry season for the three Podocnemis sampling locations.

Figure III 1. Species and sampling locations for turtle analyzed in the study. In red localities were P. vogli come from.

103 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Table III 1. Turtle species analyzed in the present study, from diverse localities in Colombia.

Sample locality Prevalence Family Species n (Department) (%) Mesoclemmys Chelidae Loríca (Cordoba) 0 dahli 3 Cravo Norte (Arauca) Podocnemis 06°14' 30,84'' N. 70° 14' 1 (100%) unifilis 1 31,05'' W Podocnemis Lorica (Cordoba) 17 lewyana 17 Podocnemididae Paz de Ariporo (Casanare) 5° 42' 18,9463'' N 71° 14' 134 134 (100%) Podocnemis 47,5309'' W vogli Puerto Carreño* (Vichada)

06°16' 067'' N. 67°49' 742'' 13 (100%) 13 W Kinosternon Kinosternidae Lorica (Cordoba) scorpioides 3 3 Trachemys

Emydidae venusta Lorica (Cordoba) 11 11 callirostris Yondó (Antioquia) 6° 48'

27,0720'' N. 74° 12' 19,69'' 5 5 Chelonoidis W carbonaria Paz de Ariporo (Casanare)

Testudinidae 5° 42' 18,9463'' N 71° 14' 1 1 47,5309'' W Cumaribo (Vichada) Chelonoidis 3° 22' 26,796'' N. 69° 30' 2 2 denticulata 48,311'' W

All individuals were captured using funnel traps, fishing nets and by hand. Blood samples were collected by puncture of the coccygeal vein and the subcarapacial sinus; up to 1% of the weight of the sampled individual was sampled. Peripheral blood smears were fixed with methanol for five minutes and stained with Giemsa 4% pH 7.2 for 45 min. Part of the blood sample was stored in an EDTA buffer (0.05 M Tris, 0.15 M NaCl, pH 8.0) or Ethanol, for further molecular analysis. The samples were kept at room temperature in the field and then refrigerated at -20 ° C in the laboratory. Capítulo 3 104

All specimens were collected under the framework permit for the collection of wild specimens of biological diversity for non-commercial purposes granted to the National

University of Colombia by the national environmental licensing authority (ANLA), resolution

0255 of March 12, 2014, and the modified resolution 1482 of November 20, 2015. The procedures performed were endorsed by the ethics committee of the Unitropico University

Foundation and the National University of Colombia. All captured individuals were released after sampling.

3.3.2 Morphological and morphometric analysis Each blood smear was examined with the 400x and 1000x objective lenses for 10 and 20 minutes, respectively using an Olympus CX41 microscope (Olympus Corporation, Tokyo,

Japan). Positive smears were analyzed for haemoparasites completely by capturing images with the Olympus DP27 digital camera and processing with the standard CellSens software. Morphometric analysis was using Image J software (Schneider et al. 2012), and we calculated the parasitaemia as the number of parasites in 10,000 erythrocytes, under the 1000x objective lense (Matta et al. 2018). Taxonomy of the haemoparasites was determined based on a comparison of morphological characteristics of the haemoparasites identified from previous reports of Haemogregarina (Siddall and Desser 1992; Telford 2009;

Soares et al. 2014; Úngari et al. 2018).

3.3.3 Molecular analysis Total DNA was extracted from samples using the phenol-chloroform method (Sambrook et al. 1989) or DNeasy (Quiagen) kit, and we measured the DNA concentration using a

NanoDrop Lite spectrophotometer (Thermo Scientific, Massachusetts, USA). PCR amplification was done using the primers HepF300 / Hep900 (Ujvari et al. 2004) to get a fragment of 600 bp of 18S rRNA from the parasites. The PCR protocol was modified from 105 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA. the original protocol as follows: denaturation at 94 ° C for 3 min, followed by 5 nonspecific cycles to ensure that the primers did not bind to other parts of the template sequence, with denaturation at 94 ° C for 30 s, annealing at 50 ºC for 45 s, and final extension at 72 º for

45 s, followed by 35 cycles with denaturation at 94 ºC for 30 s, annealing at 60 ºC for 45 s and final extension at 72 ºC for 1 min, followed by cycles of final extension at 72 ºC for 10 min. A sample that previously had been determined to be positive for Haemogregarina was used as positive control. For negative controls it was used a sample that previously had been determined to be negative for Haemogregarina. For blank, PCR was carried out without a DNA sample, only with nuclease-free water. All amplification products were evaluated by running 3 uL of the PCR on a 2% gel agarose (with EtBr?), and imaged using ultraviolet transilluminator. Amplified products were precipitated with ammonium acetate and 95% ethanol.

Sequences were edited using Sequencer v4.1.4. Each sequence was manually reviewed to verify the existence or absence of double peaks, and then aligned using ClustalW (Larkin et al. 2007) implemented in MEGA v7.0.9 (Kumar et al. 2016). The final alignment included

108 sequences of 595 nucleotides. 32 sequences previously published in Genbank of the

Hepatozoon, Hemolivia, Karyolysus and Dactylosoma genera were used. Adelina dimidiate was used as an outgroup.

Phylogeny using Bayesian inference in MrBayes version 3.2.6 through the CIPRES

Science Gateway V.3.3. The general time-reversible model was used including invariant sites and variation among sites (GTR + I + G), calculated in jModelTest 2.1.1 (Darriba et al.

2012) . The analysis was conducted with independent runs of 5 X 106 generations with six chains, sampling every 100 generations. Graphically convergence was assessed using

Tracer and phylogeny was visualized using FigTree v1.4.4. Maximum Likelihood (ML) Capítulo 3 106 analysis was done using IQTREE. Support for nodes was estimated using the bootstrap technique with 100 replicates.

Divergence was obtained within and between the different groups of sequences using the

Maximum Composite Likelihood model of substitution, implemented in the program MEGA v7.0.9 (Kumar et al. 2016).

3.4 Results

The 148 individuals infected with morphotypes belonging to the genus Haemogregarina corresponded to turtles of the genus Podocnemis, species P. unifilis and P. vogli. The general prevalence of infection was 76% (148/190) (Table III 1). All individuals were free- living. We obtained peripheral blood smears for 109 individuals, for which the associated morphology was evaluated.

3.4.1 Morphological and morphometric analyses In P. vogli all the parasitic forms identified in the individuals analyzed were found to be infecting mature erythrocytes. In the peripheral blood smears, meronts, trophozoites, and immature and mature gametocytes were identified. Based on the morphological characteristics identified in the gametocytes as described by Telford (2009), three morphotypes were identified. Morphotype 1 was found in 98% (n = 107/109) of the individuals. Morphotype 2 in 91% (n = 100/109) and morphotype 3 in 52% (n = 57/109) of these. The intensity of the infection (parasitemia) differed between the morphotypes found;

0.28 ± 0.26% (0.00 - 1.56%), 0.07 ± 0.08% (0.00 -0, 44%) and 0.01 ± 0.01% (0.00 - 0.07%), respectively. Simple infections of each morphotype in six individuals for the morphotype 107 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA. one and two individuals for the morphotype two were identified. The isolated sequences of this morphotype are shown in Table III 2.

Morphotype 1. Present in Podocnemis vogli

The immature stages are characterized by the absence of cytoplasm inside the body of the parasite giving the appearance of a "waning moon" (Fig. III 2 A). In mature gametocytes, the acytoplasmic space is reduced until it disappears (Fig. III 2. B-D). All the observed forms have the nucleus located in the periphery of the parasite. In the host cell, gametocytes cause polar displacement of the nucleus of the infected cell. All the forms observed at no time had contact with the nucleus or the membrane of the cell. The dimensions of the parasite obtained total an area of 48.36 ± 12.31 μm2, a length of 11.87 ± 1.76 μm and a width of 4.37 ± 0.71 μm.

Morphotype 2. Present in Podocnemis vogli

The mature and immature gametocytes are characterized by their small nucleus that is located towards the ends of the parasite and by agranular cytoplasm. The cytoplasm is located in the polar form in the host cell, generating the displacement of the nucleus. Like morphotype 1, none of the observed forms had contact with the nucleus or the red cell membrane (Fig. III 2 E-H). The measurements of the parasite area were 53.29 ± 9.53 μm2, length of 12,12 ± 0,99 μm, and the width was 4,97 ± 0,65 μm.

Morphotype 3. Present in Podocnemis vogli

A less prevalent morphotype was found in only 33 of the individuals sampled from the department of Casanare. Unlike the previously mentioned morphotypes, the cytoplasm was granular. The back part of the parasite was characterized by its pink hue and the absence of granules. The nucleus of the cell was displaced towards the poles, and in some cells, Capítulo 3 108 the nucleus was completely expelled, besides generating marked hypertrophy (Fig. III 2 I-

L). The size of the area was 123 ± 17.95 μm2, length of 18.2 ± 1.09 μm and width of 7.9 ±

0.94 μm.

Haemogregarina present in Podocnemis unifilis.

The gametocytes identified in this species were characterized by an absence of deformity of the infected erythrocytes. All the gametocytes observed were infecting mature erythrocytes. In these cells, the parasitic forms were located towards the pole. The gametocytes were characterized by having agranular basophilic cytoplasm; the nucleus was located towards the periphery of the parasite, occupying one third of the body of the parasite. The parasitaemia found in this individual was 0.46% (Fig III 2; M-P).

109 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Figure III 2. Haemogregarina sp. identified in Podocnemis vogli and P. unifilis . Haemogregarina sp. In P. vogli (A) Meront (B-I) Morphotype 1. (B-F) Young gamont (G-I) Mature gamont. (J-L) Morphotype 2. Mature gamont. Arrow double: Acytoplasmic space; double triangle: nucleus. Black arrows: cytoplasm pink hue. Haemogregarina sp. In P. unifilis (M-P) Mature gamont. Double triangle: nucleus. Bar 10 µm

3.4.2 Genetic distances and phylogenetic relationships In total, 121 individuals were sequenced; the maximum length of the sequences was 595 bp. Based on this information, 45 sequences were not used for the analysis, since they had double peaks in different positions. The Bayesian inference analysis and maximum likelihood used in our study allowed us the separation by the different taxa: Hepatozoon,

Hemolivia and Haemogregarina (Fig III 3-4.) with the marker used. Based on our analyses,

Haemogregarina can be divided into two differentiated clades according to their geographic Capítulo 3 110 location. In clade A, the sequences reported in the Old World are found as an ancestral group of all Haemogregarina. In clades B, C and D are the Haemogregarina reported for

South America. The sequence reported for Dactylosoma ranarum is a sister group of

Haemogregarina reported for South America. The only sequence used of Karyolysus belonged in the same clade as Hepatozoon and Hemolivia. In the case of Hepatoozoon, we obtained two clades; the first corresponded to species known for mammals (clade G), while in the other clade (clade H) all the sequences described in herpes fell into Hemolivia, a group that is a sister group of Hepatozoon (clade I). In the case of the three sequences reported as Hemogregarina Podocnemis these were isolated from Podocnemis unifilis in

Brazil. Despite the great morphological similarities with the morphotypes found in this study, the sequences grouped outside the clade of the sequences that we obtained. The distance of Haemogregarina podocnemis (clade B) from the two clades obtained in our study is

0.032 (clade C) and 0.028 (clade D). 111 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Figure III 3. A Maximum likelihood estimation hypothesis of Haemogregarina parasites based on partial sequences of 18S RNA gene (108 sequences and 595 bp). Adelina dimidiata genus used as outgroup. Capítulo 3 112

Figure III 4. A Bayesian phylogenetic hypothesis of Haemogregarina parasites based on partial sequences of 18S RNA gene (108 sequences and 595 bp). The values above branches are posterior probabilities). Adelina dimidiata was used as outgroup.

113 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Redundant sequences were not included, obtaining a total of 14 unique sequences of

Haemogregarina infecting P. vogli (Table III 2), and these showed a genetic distance between 0.002 and 0.014. For a single sequence obtained from Podocnemis unifilis, a distance was obtained with the sequences identified in P. vogli of between 0.009 and 0.021.

The sequences Pod_vog_Haemog_001 (n = 62), 002-003-004-006-007-008 and 10, grouped in clade C, with a genetic distance intragroups of 0.011. The sequence

Pod_vog_Haemog_005-009-011-012-013-014 were grouped in clade D and differed from each other by 0.007. The distance obtained between these two clades was 0.022. In the case of the sequence Pod_vog_Haemog_004, its location in the phylogenetic tree was the only one that varied according to the method used; for Bayesian inference, it was located as the ancestral sequence of clade C, while, with maximum likelihood, it was located as the ancestral sequence of the clade D.

Table III 2. Parasitaemia and sequence of each sample analyzed of Podocnemis vogli of department Vichada and Casanare-Colombia.

PARASITAEMIA ID SAMPLE SEQUENCE MORPHOTYPE 1 MORPHOTYPE 2 MORPHOTYPE 3 PV5 Pod_vog_haemog_001 0.61 0.07 0 PV4 Pod_vog_haemog_001 0.15 0.03 0 141 Pod_vog_haemog_001 0.12 0.02 0 140 Pod_vog_haemog_001 0.15 0.09 0 139 Pod_vog_haemog_001 0.04 0.02 0.02 137 Pod_vog_haemog_001 0.3 0.06 0 135 Pod_vog_haemog_001 0.13 0.02 0 133 Pod_vog_haemog_001 0.07 0.06 0 131 Pod_vog_haemog_001 0.07 0.01 0 130 Pod_vog_haemog_001 0 0.03 0 127 Pod_vog_haemog_001 0.34 0.11 0.01 126 Pod_vog_haemog_001 0.2 0.05 0.01 125 Pod_vog_haemog_001 0.14 0.03 0.02 122 Pod_vog_haemog_001 0.62 0.15 0.01 121 Pod_vog_haemog_001 0.17 0.01 0.02 120 Pod_vog_haemog_001 0.23 0.01 0.01 118 Pod_vog_haemog_001 0.16 0 0.02 115 Pod_vog_haemog_001 0.14 0 0.02 113 Pod_vog_haemog_001 0.21 0 0 110 Pod_vog_haemog_001 0.14 0.02 0 109 Pod_vog_haemog_001 0.15 0.1 0 107 Pod_vog_haemog_001 0.54 0.35 0 106 Pod_vog_haemog_001 0.23 0.01 0.02 104 Pod_vog_haemog_001 0.21 0.04 0 Capítulo 3 114

102 Pod_vog_haemog_001 0.17 0.09 0.01 101 Pod_vog_haemog_001 0.69 0.22 0.02 100 Pod_vog_haemog_001 0.33 0.01 0.01 98 Pod_vog_haemog_001 0.17 0.01 0.01 96 Pod_vog_haemog_001 0.1 0.04 0.01 94 Pod_vog_haemog_001 0.06 0.01 0 93 Pod_vog_haemog_001 0.06 0.03 0.01 92 Pod_vog_haemog_001 0.14 0.06 0.01 91 Pod_vog_haemog_001 0.23 0.15 0 85 Pod_vog_haemog_001 0.8 0.08 0 83 Pod_vog_haemog_001 0.64 0.11 0.06 79 Pod_vog_haemog_001 0.58 0.01 0.01 78 Pod_vog_haemog_001 0.5 0.08 0.07 76 Pod_vog_haemog_001 0.49 0.09 0.01 75 Pod_vog_haemog_001 0.22 0.04 0 74 Pod_vog_haemog_001 0.1 0.01 0 73 Pod_vog_haemog_001 0.17 0.06 0.01 71 Pod_vog_haemog_001 0.09 0.01 0.01 70 Pod_vog_haemog_001 0.08 0.01 0.01 69 Pod_vog_haemog_001 0.8 0.1 0.03 68 Pod_vog_haemog_001 0.65 0.11 0 67 Pod_vog_haemog_001 0.19 0.05 0 62 Pod_vog_haemog_001 0.03 0 0 58 Pod_vog_haemog_001 0.29 0.03 0 57 Pod_vog_haemog_001 0.13 0.02 0.01 52 Pod_vog_haemog_001 0.03 0.01 0 49 Pod_vog_haemog_001 0.04 0.04 0.05 45 Pod_vog_haemog_001 0.19 0.05 0.02 43 Pod_vog_haemog_001 0.27 0.2 0.03 35 Pod_vog_haemog_001 0.14 0.08 0.02 24 Pod_vog_haemog_001 0.01 0 0 22 Pod_vog_haemog_001 0.08 0.02 0 21 Pod_vog_haemog_001 0.09 0.07 0 19 Pod_vog_haemog_001 0.37 0.12 0.01 11 Pod_vog_haemog_001 0.17 0.04 0.01 4 Pod_vog_haemog_001 0.82 0.37 0.02 2 Pod_vog_haemog_001 0.42 0.09 0.02 142 Pod_vog_haemog_002 0.05 0.01 0 136 Pod_vog_haemog_003 0.37 0.16 0.01 132 Pod_vog_haemog_004 0.16 0.09 0.04 128 Pod_vog_haemog_005 0.05 0.01 0 124 Pod_vog_haemog_006 0.09 0.01 0.01 112 Pod_vog_haemog_007 0.49 0 0 88 Pod_vog_haemog_008 0.24 0.15 0.01 86 Pod_vog_haemog_009 0.15 0.01 0.02 82 Pod_vog_haemog_010 1.05 0.44 0 61 Pod_vog_haemog_011 0 0.01 0 59 Pod_vog_haemog_012 ------34 Pod_vog_haemog_013 0,07 0,03 0 31 Pod_vog_haemog_014 0,05 0,03 0

3.5 Discussion

115 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

This is the first report of haemoparasites in individuals of the free-living species of P. vogli and P. unifilis in Colombia. With relation to the microorganisms previously described in

Colombia from P. vogli to date, only the presence of nematodes such as Orientatractis leiperi (Buckley 1969) or Paraorientatractis colombiaensis (Gibbons et al. 1995) has been published. The prevalence of 89% found for the genus Haemogregarina in podocnemids agrees with the value reported in Podocnemis unifilis in Brazil of 98% (n = 72) (Soares et al. 2014) and of 94.73% (n = 36) (Úngari et al. 2018) and it is superior to the prevalence found in Podocnemis expansa of 78% (n = 96) in Peru (Pineda-Catalan et al. 2013) and

66% (n = 75) in Brazil (Picelli et al. 2015). The absence of a detectable infection in the species Podocnemis lewyana, Mesoclemmys dahli, and Trachemys venusta callirostris could be associated with (1) the distribution of these species or (2) absence of the vector implicated in the transmission; M. dahli and T. venusta callirostris are characterized as inhabiting shallow waters that may be marshy or muddy (Rueda-Almonacid et al. 2007), however, new studies related with the habitat and vector, should be developed, to evaluate these hypotheses. Our results agree with that reported by Úngari et al., (2018) who also did not find infection by Haemogregarina in K. scorpioides, nor in two species of the genus

Trachemys, T. dorbigni and T. elegans, nor in Chelonoidis sp. Soares et al. (2017) report the presence of the genus Hepatozoon, in K. scorpioides through molecular analysis.

Although P. vogli has a relatively narrow distribution, it is sympatric with P. unifilis and P. expansa (Rueda-Almonacid et al. 2007), which could explain the common presence of this genus of haemoparasites in these three species. To the date, the vector associated with the transmission of Haemogregarine are the leeches (Siddall and Desser 1993; Siddall

1995); and these organisms were found in six out the total turtles sampled, allows us to hypothesize that leeches might be responsible for the transmission of Haemogregarines in Capítulo 3 116 our localities. Further studies focused on characterizing potential vectors in the zone, are necessary.

The high prevalence of this infection in turtles is not surprising since (1.) previously, long- term follow-up studies have identified the presence of parasites for long periods in peripheral blood, such as Hepatozoon for Caiman yacare, Boa constrictor amarali and

Hydrodynastes gigas (Santos et al. 2005; Viana, et al. 2010) and Hemogregarina sp. in

Cnemidophorus arubensis (Siddall and Desser 1991) and the presence of Hemolivia mauritanica in Testudo marginata for a period ranging from 1 to 8 years (Široký, et al. 2004);

(2) turtles were present in high densities, living hundreds in the sampling areas, besides the habitats in which these turtles live are bodies of lentic water that could facilitate the infection. Besides, if the vector were leeches (which also share this habitat) they can be detached from a host and then adhere to the new one, which increases the possibility of transmission among the individuals that live there (3.) On the other hand, it has been reported a low pathogenicity of haemoparasites of the genera Haemogregarina and

Hepatozoon, that could also explain their presence in high prevalences.

Parasites such as Hepatozoon and Haemogregarina have been considered non- pathogenic, which could also be associated with the high prevalence of infection. Damas-

Moreira et al. (2014) report that parasites such as Hepatozoon generated limited effects in the lizard Podarcis vaucheri. Davis and Sterrett (2011) report that infection by

Haemogregarina sp. in Trachemys scripta, Chrysemys picta and Sternotherus odoratus have no obvious effects on the health status of these individuals. However, studies conducted by Bouma et al. (2007) report that Tiliqua rugosa lizards infected by Hemolivia mariae decrease their range of distribution, compared with non-infected hosts. For a long time similar effects were speculated for blood parasites in wild birds and recently a series of reports show that parasites such as Haemoproteus can be pathogenic and cause 117 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA. deleterious effects on their host (Earle et al. 1993; Ilgūnas et al. 2016; Ortiz-Catedral et al.

2019) which is why we speculate that studies that follow infected reptiles probably might conclude with similar results.

Regarding the intensity of parasitemia, the highest values found in this study was in

Podocnemis vogli with 1.49% (morphotype one and two); and in P. unifilis the parasitaemia was 0.46%. These values were lower than reported for Haemogregarina podocnemis in P. unifilis, which the parasitaemia was 2.96%, the highest known value in free-living turtles is

4.03% and for captive animals 1.53% (Úngari et al. 2018). In P. unifilis, Soares et al. (2014) reported an average of parasitemia of 6%. Our results are also lower than those reported for P. expansa in Brazil, with a prevalence of 3%. It is worth noting that in our study the parasitaemia was calculated of the morphotype three by separated; while that in other studies the parasitaemia was obtained without differentiating between morphotypes. For example, Úngari et al. (2018) in the description of Haemogregarina podocnemis considered the three morphotypes as only one species and thus determine parasitaemia based on that.

The morphotypes reported in P. vogli share morphological characteristics with the gamonts identified by Soares et al. (2014) and those of the species Haemogregarina podocnemis, isolated from P. unífilis in Brazil (Úngari et al. 2018). In both studies, different morphotypes were seen; however, these authors grouped them as micro- and macrogamonts. At this point, it is important to highlight, the absence of sexual dimorphism for Haemogregarina species (Telford 2009). Moreover, group different morphotypes as only one species is risky, considering that these morphotypes could be the result of a coinfection. It is necessary to emphasize that the morphotypes reported in the studies carried out in Brazil are characterized by the presence of a parasitophorous vacuole, which, in the case of Soares Capítulo 3 118 et al. (2014), was present in all the observed morphotypes. This feature was not present in the gamonts described in our study.

Given that the same molecular sequence (Pod_vog_Haemog_001) was obtained in 61 individuals (that showed the morphotype 1), and five with morphotype 2 (Table III 2), and all of them showed single infection microscopically, we can suggest morphotypes 1 and 2 could represent morphological variants of the same parasite. Unfortunately, morphotype 3 was never found in a simple infection, and the sequences obtained from the organisms that carried it was equal to the other sequences found for infections of morphotypes 1 and 2; other methodologies such as cloning, design of new primers or new molecular markers should help to solve this problem.

The genetic distance between clades C and D (0.022) is less than the distance of these two with the Haemogregarina sequences described in the Old World (0.111 and 0.107, respectively) and it is even less than the distance obtained with Haemogregarina podocnemis, described in Brazil. The genetic distance between the morphotypes identified in the study and Haemogregarina podocnemis, despite the parasitic forms observed in blood being indistinguishable, suggests that they could be cryptic species, taking into account the genetic distance between the sequences obtained in our study and the reported in Brazil. In the results of this study, no geographical differentiation of the identified sequences was observed (Table 2). Several of the sequences obtained were different in one or a few nucleotides resulting in low genetic distances, these results are similar to reported from Leucocytoozon infecting blue tits, where two new sequences were identified that differ each in 0,21% from the sequence more closest reported in MalAvi, or in

Plasmodium, which the genetic distance between sequence obtained was between 0.23% and 4.32% (Schumm et al. 2019). 119 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

The distance obtained between the sequences grouped in clade C was 0.011 and it was equal to the distance obtained for Hemolivia sp. (n = 4) (DvořáKová et al. 2015), while the distance obtained between sequences in clade D (0.007) is higher than that reported between Haemogregarina balli and Haemogregarina stepanowi (0.0017) (DvořáKová et al.

2014). Authors such as O'Dwyer et al. (2013) describe new species of Hepatozoon infecting snakes, based on the differentiation of one or two base pairs; however, this approach could be erroneous, bearing in mind that this variation can be the result of the presence of multiple copies of the fragment 18s RNA, taking into account that in the ribosomal rRNA genes can have multiples copies these differ minimally between them (Le Blancq et al. 1997). In our analyses, as previously mentioned, 45 sequences were obtained in which multiple peaks were seen in some positions, which could be associated with the presence of these multiple copies of 18s RNAr that can differ in a few bases in the part variable of the sequence. Our results show that intraspecific differentiation for Haemogregarina is complex, as reported by Maia (2015) for the genus Hepatozoon in the species H. chalcides, H. eumeces, H. atlantolacerta, H. timon and H. podarcis, where these multiple haplotypes constitute a completely new genetic sequence.

Taking into account the lack of characteristics of sexual dimorphism in the Haemogregarina, we disagree with the reason exposed by Úngari et al., (2018) for join all morphotyphes as a species. In our case, the absence of the third morphotype in 52 of 147 individuals analyzed, and specifically their absence in the town of Puerto Carreño-Vichada, lead us to think that the morphotype three does not form part of the same species. Despite this, in co- infected organisms (Table III 2), it was not possible to isolate a different sequence from that shown in morphotypes 1 and 2. The possible explanation may lie in (1.) the low parasitaemia of this morphotype; or (2.) the primers could not match correctly to sequences of this morphotype (a similar situation was recently reported in a detection failure of Capítulo 3 120

Leucocytozoon in birds (Lotta et al. 2019). The match of primers used in this study and others reported in the literature shows in the Supplementary figure III 1.

The presence of the third morphotype in P. unifilis in Brazil and in P. vogli in Colombia, could represent the same parasite associated with the range of distribution of these turtle species. In our study we did not find a geographical separation of the identified sequences, sharing the Pod-vog-Haemo_001 sequence between the two locations sampled, despite a large geographical distance (approximately 1068 km) (Table III 2). However, these two localities: Puerto Carreño and Paz de Ariporo, share ecosystem characteristics such as being both floodplains, bathed by the Orinoco, Guayabero and Guaviare (Forero-Medina et al. 2014), and their aquatic tributaries, which would allow the movement of individuals between them and the circulation of parasites between these geographic areas, however, until now, the authors have no found any migration patterns described for P. vogli.

The genetic distance between the sequence of the morphotype identified in P. unifilis with those found in P. vogli did not exceed 0.010, despite the morphological differences. This underlining the need to group both the information obtained by microscopy and that obtained from the phylogenetic analysis, when describing species.

Although to date, the 18s rRNA gene is the only available marker for the identification of the different genera belonging to the Adeleorina suborder, this gene can have multiple copies that can be identified; also in Plasmodium, have been reported different types of rRNA according to the phase of the lifecycle (Sexual in the vector; Asexual in de host vertebrate); and this gene shows highly conservative sites and other variables. However, as observed in the analysis of Bayesian inference and maximum likelihood (Fig. III 3 and

4), the values obtained in the most recent nodes are decreasing. Also, it should be taken into account that in this group of haemoparasites using this marker, the phylogenetic 121 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA. relationships are not fully resolved, so reassignment of species even among genera continues to be common (Cook et al. 2014).

Regarding the presence of multiple peaks in the analyzed sequences, this may be due to the presence of co-infections in the same sample, which are not distinguishable by small variations in the sequences obtained from the gene; Maia (2015) reported the presence of coinfections in reptiles infected with Hepatozoon, where different associated haplotypes were reported. Co-infections are common in wildlife and have been a challenge for researchers trying to devise methodological approaches that allow their correct identification and define the actual diversity of blood parasites (Falk et al. 2015; Pacheco et al. 2018). Other possible cause of multiple peaks in the sequence can be the presence of multiples copies of the gene. For the 18s rRNA there are reports which show multiple copies that differ minimally, this could contribute erroneously to the description of species under the presumption that, since it is a fairly conservative marker small variations are indicative of new species. For example, Rooney (2004) found that Plasmodium berghei sequences differ between 76 and 154 nucleotides.

Taking into account the new challenges and difficulties related to the identification of species based on the analysis of the 18s rRNA fragment, it is necessary to develop new approaches to try to solve this problem such as 1. to standardize different methodologies to obtain a fragment greater than 600 bp, taking into account the presence of multiple copies or the slow evolutionary rate of these markers, 2. to identify new molecular markers, whether mitochondrial, apicoplast, or other nuclear markers that, together with the information already available, allows better elucidation of phylogenetic relationships, as well as internal variations of species that may arise. Capítulo 3 139

3.6 Conclusion and perspectives

This is the first study that demonstrates the presence of Haemogregarina spp., in

Podocnemis vogli, expanding the geographic and host range for this genus, and reporting

15 new sequences of these parasites in Podocnemis vogli and P. unifilis. This information can be useful in wildlife conservation and relocation programs. Based on our results, the

18s rRNA molecular marker has shown a utility to differentiate parasites at the genus level, but its usefulness is limited in more derived taxa. Therefore, we suggest a search for new markers that contribute to an improvement in the resolution of phylogenetic relationships of parasites of the suborder Adeleorina, as well as, to intra-specific differentiation. It also is very important to design studies that allow the identification of vectors allowing a better characterization of life cycles.

3.7 Acknowledgements

This study was supported by the Administrative Department of Science, Technology and

Innovation COLCIENCIAS FP2018-18-44842 and the inter-institutional agreement between Unitrópico and Parex Resources "Huella Galápaga", the University Foundation -

Unitrópico. The authors thank the support of the members of the GERPH research group, especially Ingrid Lotta, for their support in the development of the analyses. Additionally, to

GINBIO research members, Parales family in the natural reserve “la Esperanza” and Rafael

Gutierrez, for their field efforts. The authors would also like to thank to Cristian Rodriguez for the images of Podocnemis vogli used in figure III 1 and graphical abstract.

Figure III 1. Species and sampling locations for turtle analyzed in the study. In red localities were P. vogli come from.

Figure III 2. Haemogregarina sp. identified in Podocnemis vogli and P. unifilis . Haemogregarina sp. In P. vogli (A) Meront (B-I) Morphotype 1. (B-F) Young gamont (G-I) Mature gamont. (J-L) Morphotype 2. Mature gamont. Arrow double: Acytoplasmic space; 140 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA. double triangle: nucleus. Black arrows: cytoplasm pink hue. Haemogregarina sp. In P. unifilis (M-P) Mature gamont. Double triangle: nucleus. Bar 10 µm

Figure III 3. A Maximum likelihood estimation hypothesis of Haemogregarina parasites based on partial sequences of 18s rRNAgene (108 sequences and 595 bp). Adelina dimidiata genus used as outgroup.

Figure III 4. A Bayesian phylogenetic hypothesis of Haemogregarina parasites based on partial sequences of 18s rRNA gene (108 sequences and 595 bp). The values above branches are posterior probabilities). Adelina dimidiata was used as outgroup

Table III 1. Turtle species analyzed in the present study, from diverse localities in Colombia.

Table III 1. Parasitaemia and sequence of each sample analyzed of Podocnemis vogli of department Vichada and Casanare-Colombia.

3.8 Reference

Adl SM, Bass D, Lane CE, et al (2019) Revisions to the Classification, Nomenclature, and Diversity of Eukaryotes. Journal of Eukaryotic Microbiology. doi: 10.1111/jeu.12691

Bouma MJ, Smallridge CJ, Bull CM, Komdeur J (2007) Susceptibility to infection by a haemogregarine parasite and the impact of infection in the Australian sleepy lizard Tiliqua rugosa. Parasitology Research 100:949–954. doi: 10.1007/s00436-006- 0379-5

Brown GP, Shilton CM, Shine R (2006) Do parasites matter? Assessing the fitness consequences of haemogregarine infection in snakes. Can J Zool 84:668–676. doi: 10.1139/z06-044

Buckley JJC (1969) On a Remarkable Oxyurid Nematode, Orientatractis leiperi n. sp., (Atractidae) from a South American Tortoise, Podocnemis vogli. J Helminthol 43:281–286. doi: 10.1017/S0022149X0000482X

Cook CA, Netherlands EC, Smit NJ (2016) Redescription, molecular characterisation and taxonomic re-evaluation of a unique African monitor lizard haemogregarine Karyolysus paradoxa (Dias, 1954) n. comb. (Karyolysidae). Parasites & Vectors 9:. doi: 10.1186/s13071-016-1600-8 Capítulo 3 124

Karyolysus paradoxa (Dias, 1954) n. comb. (Karyolysidae). Parasites & Vectors 9:. doi: 10.1186/s13071-016-1600-8

Cook CA, Smit NJ, Davies AJ (2010) Hemoproteids (Apicomplexa: Haemoproteidae) from South African Tortoises (Cryptodira: Testudinidae). Journal of Parasitology 96:1168–1172. doi: 10.1645/GE-2527.1

Dalrymple BP (1990) Cloning and characterization of the rRNA genes and flanking regions from Babesia bovis: use of the genes as strain discriminating probes. Molecular and Biochemical Parasitology 43:117–124. doi: 10.1016/0166-6851(90)90136-A

Damas-Moreira I, Harris DJ, Rosado D, et al (2014) Consequences of haemogregarine infection on the escape distance in the lacertid lizard, Podarcis vaucheri. Acta Herpetologica Vol 9:119-123 Pages. doi: 10.13128/acta_herpetol-13740

Darriba D, Taboada GL, Doallo R, Posada D (2012) jModelTest 2: more models, new heuristics and parallel computing. Nature Methods 9:772–772. doi: 10.1038/nmeth.2109

Davis AK, Sterrett SC (2011) Prevalence of Haemogregarine Parasites in Three Freshwater Turtle Species in a Population in Northeast Georgia, USA. International J of Zoological Research 7:156–163. doi: 10.3923/ijzr.2011.156.163 de Oliveira JP, André MR, Alves Júnior JR, et al (2018) Molecular detection of hemogregarines and haemosporidians in Brazilian free-living testudines. International Journal for Parasitology: Parasites and Wildlife 7:75–84. doi: 10.1016/j.ijppaw.2018.01.008

DvořáKová N, KvičErová J, Hostovský M, šIroký P (2015) Haemogregarines of freshwater turtles from Southeast Asia with a description of Haemogregarina sacaliae sp. n. and a redescription of Haemogregarina pellegrini Laveran and Pettit, 1910. Parasitology 142:816–826. doi: 10.1017/S0031182014001930

DvořáKová N, KvičErová J, PapoušEk I, et al (2014) Haemogregarines from western Palaearctic freshwater turtles (genera Emys, Mauremys) are conspecific with Haemogregarina stepanowi Danilewsky, 1885. Parasitology 141:522–530. doi: 10.1017/S0031182013001820

Earle RA, Bastianello SS, Bennett GF, Krecek RC (1993) Histopathology and morphology of the tissue stages of Haemoproteus columbae causing mortality in Columbiformes. Avian Pathology 22:67–80. doi: 10.1080/03079459308418901

Falk BG, Glor RE, Perkins SL (2015) Clonal reproduction shapes evolution in the lizard malaria parasite Plasmodium floridense. Evolution 69:1584–1596. doi: 10.1111/evo.12683

Forero-Medina G, Yusti-Muñoz AP, Castaño-Mora OV (2014) Geographic Distribution of the Tortoises and Freshwater Turtles of Colombia and their Representation in the Protected Area Network. Acta Biológica Colombiana 19:415. doi: 10.15446/abc.v19n3.42219 125 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Gaffney ES, Meylan PA, Wood RC, et al (2011) Evolution of the side-necked turtles: the family Podocnemididae. Bulletin of the American Museum of natural History

García-Longoria L, Møller AP, Balbontín J, et al (2015) Do malaria parasites manipulate the escape behaviour of their avian hosts? An experimental study. Parasitol Res 114:4493–4501. doi: 10.1007/s00436-015-4693-7

Gibbons LM, Khalil LF, Marinkelle CJ (1995) A new nematode genus, Podocnematractis, for Atractis ortleppi Thapar, 1925 (Cosmocercoidea: Atractidae) and the description of another new species, P. colombiaensis from turtles Podocnemis spp. in Colombia. Syst Parasitol 30:47–56. doi: 10.1007/BF00009244

Gutierrez G (2016) PARÁSITOS SANGUÍNEOS EN ALGUNOS REPTILES DE LA LOCALIDAD DE YONDÓ, ANTIOQUIA, COLOMBIA. Universidad Nacional de Colombia

Hoffmann M, Hilton-Taylor C, Angulo A, et al (2010) The Impact of Conservation on the Status of the World’s Vertebrates. Science 330:1503–1509. doi: 10.1126/science.1194442

Ilgūnas M, Bukauskaitė D, Palinauskas V, et al (2016) Mortality and pathology in birds due to Plasmodium (Giovannolaia) homocircumflexum infection, with emphasis on the exoerythrocytic development of avian malaria parasites. Malar J 15:256. doi: 10.1186/s12936-016-1310-x

Kibe MK, ole-MoiYoi OK, Nene V, et al (1994) Evidence for two single copy units in Theileria parva ribosomal RNA genes. Molecular and Biochemical Parasitology 66:249–259. doi: 10.1016/0166-6851(94)90152-X

Kumar S, Stecher G, Tamura K (2016) MEGA7: Molecular Evolutionary Genetics Analysis Version 7.0 for Bigger Datasets. Molecular Biology and Evolution 33:1870–1874. doi: 10.1093/molbev/msw054

Lainson R, Naiff RD (1998) Haemoproteus (Apicomplexa: Haemoproteidae) of Tortoises and Turtles. Proceedings: Biological Sciences 265:941–949

Laird M (1950) Haemogregarina tuatarae sp. n., from the New Zealand Rhynchocephalian Sphenodon punctatus (Gray). Proceedings of the Zoological Society of London 120:529–533. doi: 10.1111/j.1096-3642.1950.tb00662.x

Larkin MA, Blackshields G, Brown NP, et al (2007) Clustal W and Clustal X version 2.0. Bioinformatics 23:2947–2948. doi: 10.1093/bioinformatics/btm404

Le Blancq SM, Khramtsov NV, Zamani F, et al (1997) Ribosomal RNA gene organization in Cryptosporidium parvum 1Note: Nucleotide sequence data reported in this paper have been submitted to the GenBankTM data base under the accession numbers AF015772, AF015773 and AF015774.1. Molecular and Biochemical Parasitology 90:463–478. doi: 10.1016/S0166-6851(97)00181-3 Capítulo 3 126

Lotta IA, Valkiūnas G, Pacheco MA, et al (2019) Disentangling Leucocytozoon parasite diversity in the neotropics: Descriptions of two new species and shortcomings of molecular diagnostics for leucocytozoids. International Journal for Parasitology: Parasites and Wildlife 9:159–173. doi: 10.1016/j.ijppaw.2019.05.002

Maia JP (2015) Diversity, infection patterns and hostparasite associations of apicomplexan parasites in reptiles. (Doctoral dissertation, Ph. D. Thesis. Universidade do Porto, Porto, Portugal).

Matta NE, González LP, Pacheco MA, et al (2018) Plasmodium parasites in reptiles from the Colombia Orinoco-Amazon basin: a re-description of Plasmodium kentropyxi Lainson R, Landau I, Paperna I, 2001 and Plasmodium carmelinoi Lainson R, Franco CM, da Matta R, 2010. Parasitology Research 117:1357–1370. doi: 10.1007/s00436-018-5815-9

McCutchan TF, Li J, McConkey GA, et al (1995) The cytoplasmic ribosomal RNAs of Plasmodium spp. Parasitol Today (Regul Ed) 11:134–138

Merino S, Moreno J, José Sanz J, Arriero E (2000) Are avian blood parasites pathogenic in the wild? A medication experiment in blue tits (Parus caeruleus ). Proceedings of the Royal Society of London Series B: Biological Sciences 267:2507–2510. doi: 10.1098/rspb.2000.1312

Morales-Betancourt MA, Lasso CA, Páez VP, Bock BC (2015) Libro rojo de reptiles de Colombia. Instituto de Investigación de Recursos Biológicos Alexander Von Humboldt, Colombia

Morand S, Krasnov BR, Littlewood DTJ (eds) (2015) Parasite diversity and diversification: evolutionary ecology meets phylogenetics. Cambridge University Press, Cambridge, United Kingdom

Morrison DA (2009) Evolution of the Apicomplexa: where are we now? Trends in Parasitology 25:375–382. doi: 10.1016/j.pt.2009.05.010

Netherlands EC, Cook CA, Smit NJ (2014) Hepatozoon species (Adeleorina: Hepatozoidae) of African bufonids, with morphological description and molecular diagnosis of Hepatozoon ixoxo sp. nov. parasitising three Amietophrynus species (Anura: Bufonidae). Parasites & Vectors 7:. doi: 10.1186/s13071-014-0552-0

Oppliger A, Célérier ML, Clobert J (1996) Physiological and behaviour changes in common lizards parasitized by haemogregarines. Parasitology 113:433–438. doi: 10.1017/S003118200008149X

Orkun Ö, Güven E (2013) A New Species of Haemoproteus from a Tortoise (Testudo graeca) in Turkey, with Remarks on Molecular Phylogenetic and Morphological Analysis. Journal of Parasitology 99:112–117. doi: 10.1645/GE-3100.1 127 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Ortiz-Catedral L, Brunton D, Stidworthy MF, et al (2019) Haemoproteus minutus is highly virulent for Australasian and South American parrots. Parasites Vectors 12:40. doi: 10.1186/s13071-018-3255-0

Ortiz-Moreno ML, Rodríguez-Pulido JA (2017) Estado del conocimiento y amenazas de la tortuga sabanera (Podocnemis vogli, Podocnemididae) en Colombia. Orinoquia 21:26. doi: 10.22579/20112629.391

Pacheco MA, Cepeda AS, Bernotienė R, et al (2018) Primers targeting mitochondrial genes of avian haemosporidians: PCR detection and differential DNA amplification of parasites belonging to different genera. International Journal for Parasitology 48:657–670. doi: 10.1016/j.ijpara.2018.02.003

Paperna I (2006) Hemolivia mauritanica (Haemogregarinidae: Apicomplexa) infection in the tortoise Testudo graeca in the Near East with data on sporogonous development in the tick vector Hyalomna aegyptium. Parasite 13:267–273. doi: 10.1051/parasite/2006134267

Picelli AM, Carvalho AV de, Viana LA, Malvasio A (2015) Prevalence and parasitemia of Haemogregarina sp. in Podocnemis expansa (Testudines: Podocnemididae) from the Brazilian Amazon. Revista Brasileira de Parasitologia Veterinária 24:191–197. doi: 10.1590/S1984-29612015033

Rooney AP (2004) Mechanisms Underlying the Evolution and Maintenance of Functionally Heterogeneous 18S rRNA Genes in Apicomplexans. Molecular Biology and Evolution 21:1704–1711. doi: 10.1093/molbev/msh178

Rossow JA, Hernandez SM, Sumner SM, et al (2013) Haemogregarine infections of three species of aquatic freshwater turtles from two sites in Costa Rica. International Journal for Parasitology: Parasites and Wildlife 2:131–135. doi: 10.1016/j.ijppaw.2013.02.003

Rueda-Almonacid, Carr, J. L., Mittermeier, R. A, Rodríguez-Mahecha, J. V., et al (2007) Las tortugas y los cocodrilianos de los países andinos del trópico. Serie de guías tropicales de campo, 6:412–423

Sambrook J, Fritch E, Maniatis T (1989) Molecular cloning: A laboratory manual, 2nd end. Cold Spring Harbour Laboratory, Cold Spring Harbor

Santos MM de V, O’Dwyer LH, Silva RJ da (2005) Seasonal variation of Hepatozoon spp. (Apicomplexa, Hepatozoidae) parasitemia from Boa constrictor amarali (Serpentes, Boidae) and Hydrodynastes gigas (Serpentes, Colubridae). Parasitol Res 97:94– 97. doi: 10.1007/s00436-005-1385-8

Schall JJ (1983) Lizard malaria: cost to vertebrate host’s reproductive success. Parasitology 87:1–6. doi: 10.1017/S0031182000052367 Capítulo 3 128

Schneider CA, Rasband WS, Eliceiri KW (2012) NIH Image to ImageJ: 25 years of image analysis. Nature Methods 9:671–675. doi: 10.1038/nmeth.2089

Schumm YR, Wecker C, Marek C, et al (2019) Blood parasites in Passeriformes in central Germany: prevalence and lineage diversity of Haemosporida (Haemoproteus , Plasmodium and Leucocytozoon) in six common songbirds. PeerJ 6:e6259. doi: 10.7717/peerj.6259

Siddall ME (1995) Phylogeny of Adeleid Blood Parasites with a Partial Systematic Revision of the Haemogregarine Complex. Journal of Eukaryotic Microbiology 42:116–125. doi: 10.1111/j.1550-7408.1995.tb01551.x

Siddall ME, Desser SS (1991) Merogonic Development of Haemogregarina balli (Apicomplexa: Adeleina: Haemogregarinidae) in the Leech Placobdella ornata (Glossiphoniidae), Its Transmission to a Chelonian Intermediate Host and Phylogenetic Implications. The Journal of Parasitology 77:426. doi: 10.2307/3283131

Siddall ME, Desser SS (1992) Prevalence and intensity of Haemogregarina balli (Apicomplexa: Adeleina: Haemogregarinidae) in three turtle species from Ontario, with observations on intraerythrocytic development. Canadian Journal of Zoology 70:123–128. doi: 10.1139/z92-018

Siddall ME, Desser SS (1993) Ultrastructure of merogonic development of Haemogregarina (sensu lato) myoxocephali (Apicomplexa: Adeleina) in the marine leech Malmiana scorpii and localization of infective stages in the salivary cells. European Journal of Protistology 29:191–201. doi: 10.1016/S0932-4739(11)80273-7

Široký P, Kamler, M, Modrý, D (2005) Prevalence of Hemolivia mauritanica (Apicomplexa: Adeleina: Haemogregarinidae) in natural populations of tortoises of the genus Testudo in the east Mediterranean region. Folia Parasitol, 52:359–361

Široký, P, Kamler, M, Modrý, D. (2004) Long-term occurrence of Hemolivia cf. mauritanica (Apicomplexa: Adeleina: Haemogregarinidae) in captive Testudo marginata (Reptilia: Testudinidae): evidence for cyclic merogony? Journal of Parasitology 90:1391–1393

Sloboda M, Kamler M, Bulantová J, et al (2007) A NEW SPECIES OF HEPATOZOON (APICOMPLEXA: ADELEORINA) FROM PYTHON REGIUS (SERPENTES: PYTHONIDAE) AND ITS EXPERIMENTAL TRANSMISSION BY A MOSQUITO VECTOR. Journal of Parasitology 93:1189–1198. doi: 10.1645/GE-1200R.1

Telford SRJr (2009) Hemoparasites of the Reptilia: color atlas and text. Taylor & Francis, Boca Raton

Ujvari B, Madsen T, Olsson M (2004) High Prevalence of Hepatozoon Spp. (Apicomplexa, Hepatozoidae) Infection in Water Pythons (Liasis fuscus) From Tropical Australia. Journal of Parasitology 90:670–672. doi: 10.1645/GE-204R

Vargas-Ramirez M, Castaño-Mora O, Fritz U (2008) Molecular phylogeny and divergence times of ancient South American and Malagasy river turtles (Testudines: Pleurodira: Podocnemididae). Organisms Diversity & Evolution 8:388–398. doi: 10.1016/j.ode.2008.10.001

Viana, LA, Paiva, F, Coutinho, M. E, Lourenço-de-Oliveira, R (2010) Hepatozoon caimani (Apicomplexa: Hepatozoidae) in wild caiman, Caiman yacare, from the Pantanal region. Journal of Parasitology, 96:83–88

Vilcins I-ME, Ujvari B, Old JM, Deane E (2009) Molecular and Morphological Description of a Hepatozoon Species in Reptiles and Their Ticks in the Northern Territory, Australia. Journal of Parasitology 95:434–442. doi: 10.1645/GE-1725.1

Zamudio N, Ramírez M (2007) Presence of Hepatozoon spp. In serpents of the center of attention and valuation of wildlife (cav) of the área metropolitana del Valle de Aburrá, Barbosa – Antioquia. Revista CES / Medicina Veterinaria y Zootecnia 2:

Capítulo 3 130

3.9 Appendix

Supplementary figure III 1. N. Nucleotide position where the primers have to generate the match with the 18S RNA sequence of Haemogregarina. Blue: Primers HEP300-HEP900. Orange: Primers HEMO1-HEMO2; Green: Primers EF-ER. The sequence used was Haemogregarina balli. Genbank HQ224959.

IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

4. Capítulo 4. Discusión general

El estudio de hemoparásitos en la herpetofauna colombiana, es un campo de investigación nuevo en nuestro país; durante el desarrollo de este estudio se generó información novedosa para el mundo como: (1) se obtuvieron las secuencias de citocromo b, para las especies P. kentropyxi y P. carmelinoi encontradas infectando Cnemidophorus cf. gramivagus y Ameiva ameiva, respectivamente, en el Departamento del Guaviare-

Colombia, descritas previamente en Brasil, unicamente con base en características morfológicas; (2) se describió la nueva especie Haemocystidium atakurum en Podocnemis vogli en los llanos orientales de Colombia, y (3) se reportó la presencia de nuevos linajes y morfotipos para el género Haemogregarina infectando Podocnemis vogli y Podocnemis unifilis en los Llanos orientales de Colombia.

Los reportes previos asociados a hemoparásitos en reptiles de vida silvestre en Colombia, se basaron exclusivamente en los caracteres morfológicos de las formas observadas en sangre (Ayala S.C et al. 1973; Ayala 1975; Ayala S.C and Spain 1976; Moreno et al. 2015;

Gutierrez 2016); Ayala y colaboradores, realizaron sus investigaciones con muestras obtenidas en las Islas de San Andrés y Providencia, el departamento del Cauca y Llanos

Orientales de Colombia. Esta última localidad abarca el departamento de Vichada, del cual provienen muestras analizadas en los capítulos dos y tres, no obstante, en el estudio realizado por Ayala et al., (1973) no fueron muestreadas tortugas, por su parte, se 132 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA. reportaron Haemogregarinas y Trypanosomas en marsupiales, y de microfilaria y haemogregarinas en Tupinambus sp.

En nuestro estudio la baja prevalencia de infección por hemosporidios, contrastó con la alta prevalencia de haemogregarinas, esto podría asociarse: (1) La mayoría de muestras analizadas corresponden a tortugas, organismos en los cuales el parásito más comúnmente encontrado son las haemogregarinas, y en los que a la fecha no se ha reportado la presencia de Plasmodium (Ayala 1978; Telford 1984); (2) En el caso de las muestras obtenidas en el departamento del Guaviare, podría presentarse un efecto de dilución relacionado con una elevada diversidad de hospederos. En Guaviare, se ha registrado un gran número de especies de fauna, específicamente, en reptiles se han registrado 67 especies de reptiles (Medina-Rangel et al. 2019). Esta diversidad, podría generar que los vectores se alimenten de hospederos vertebrados donde la infección no sea viable, o que estos vectores deban competir con otras especies no vectores por la alimentación.

En relación con la ausencia de hemoparásitos en tortugas terrestres (Ch. carbonaria y Ch. denticulata), nuestros resultados concuerdan con los reportados por Úngari et al., (2018) para Chelonoidis sp; sin embargo, en los dos casos el número de individuos muestreados fue limitado, por lo cual es necesario realizar nuevos estudios donde se aumente el número de individuos analizados para estas especies. Diversos estudios han reportado la presencia de hemoparásitos en el género Chelonoidis: Haemogregarina sp. infectando a

Ch. denticulata en Perú (Batalla L. et al. 2015) y a Chelonoidis chilensis en Paraguay

(Pedrozo Prieto et al. 2016), así como de Haemosporidios infectando Ch. carbonaria y Ch. denticulata en Brasil (Martinele et al. 2016). A nivel mundial, se ha reportado la presencia de Haemosporidios en Testudo graeca y Testudo horsfieldii (Javanbakht et al. 2015), y de Capítulo 4 133

Hemolivia párvula en Kinixys zombensis (Cook et al. 2015), especies de tortugas terrestres.

De acuerdo a la revisión literaria realizada, los reportes de hemoparásitos pertenecientes al género Haemogregarinas, son más comunes en tortugas acuáticas (Davis and Sterrett

2011; DvořáKová et al. 2014, 2015; Úngari et al. 2018) que en terrestres, esto podría asociarse con la facilidad de estar en contacto con los vectores implicados a la fecha en la transmisión de este hemoparásito, las sanguijuelas (Siddall and Desser 1993).

La ausencia de hemoparásitos en M. dahli, P. lewyana, K. scorpioides y T. venusta callirostris, puede relacionarse (1) con la distribución de estas poblaciones transandinas o

(2) la ausencia de los vectores implicados, sin embargo, estudios enfocados en evaluar estas características deben ser desarrollados para confirmar estas hipótesis, teniendo en cuenta que estos géneros de tortugas se ha reportado la presencia de hemoparásitos. M. dahli y P. lewyana son especies endémicas, lo cual no permite comparar los resultados obtenidos con estudios realizados previamente, al ser este el primer estudio enfocado en identificar los hemoparásitos que pueden infectarlas; no obstante, en estos dos géneros se ha reportado la presencia de Haemogregarinas, en las especies Podocnemis unifilis

(Soares et al. 2014; Úngari et al. 2018), Podocnemis expansa (Picelli et al. 2015),

Podocnemis vogli (Capítulo 3) y Mesoclemmys vanderhaegei (Goes et al. 2018), al igual que en el caso del género Trachemys se ha reportado la presencia de este género de hemoparásitos, en Trachemys scripta . En el caso de K. scorpioides, para esta especie se ha reportado la presencia de Haemogregarinas en Brasil (Rossow et al. 2013).

En cuanto a la identificación de hemoparásitos en herpetos, hasta hace algunas décadas este proceso se basó exclusivamente en análisis microscópico, describiéndose especies con base en las características de los parásitos en los extendidos sanguíneos de los hospederos vertebrados; sin embargo, a pesar que esta metodología es considerada el gol estándar, en la parasitología, nuestros resultados apoyan la necesidad de complementar 134 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA. la información obtenida por microscopía, con análisis moleculares dirigidos a marcadores moleculares como cyt b y 18S rRNA, conocido como taxonomía integrativa.

La descripción de nuevas especies para los géneros pertenecientes al suborden

Adeleorina, (como las hemogregarinas) no puede realizarse únicamente con base en caracteres morfológicos de los estadios observados en sangre periférica, denominados como gamontes (Telford 2009); con esta información en la mayoría de los casos la diferenciación puede realizarse hasta género, con base en caracteres, como la ausencia/presencia de merogonia eritrocitaria o los efectos que genere la infección en la célula hospedera (por ejemplo, la lisis del núcleo celular), sin embargo, la ausencia de caracteres diagnósticos para estos géneros, dificulta su caracterización. El análisis conjunto de análisis moleculares y la caracterización del ciclo de vida en el vector, es información que complementa la caracterización morfológica, conllevando a la reclasificación de especies descritas con base en características morfológicas en diferentes géneros, como Haemogregarina fitzsimonsi dentro del género Hepatozoon

(Cook et al. 2014), o Hepatozoon paradoxa como Karyolysus paradoxa (Cook et al. 2016)

En cuanto al marcador molecular utilizado para la caracterización de los hemoparásitos pertenecientes al suborden Adeleorina, el 18S rRNA, en nuestro estudio utilizamos los primers HEP 300-HEP 900 (Ujvari et al. 2004), los cuales anclan en una zona conservada de la secuencia, que nos permite amplificar secuencias de géneros como Haemogregarina,

Hepatozoon y Hemolivia; a pesar que el fragmento obtenido de aproximadamente 600 pb, corresponde a una región variable que nos permite diferenciar entre géneros y ha sido utilizado para describir especies. En la literatura se han reportado otros primers para la amplificación de fragmentos de 18S rRNA, sin embargo, es necesario evaluar los resultados que son obtenidos con estos, por ejemplo, para los primers propuestos por Capítulo 4 135

Wozniak et al., (1994), Perkins y Mathew (1999) demostraron que realmente no eran útiles para el diagnóstico de Haemogregarinas, teniendo en cuenta que amplifica fragmentos de

18S rRNA de hospederos vertebrados y de otros parásitos como Plasmodium. En cuanto a los primers HEMO 1 y HEMO 2 (Perkins and Keller 2001), no se obtuvieron resultados para las muestras amplificadas, esto puede asociarse a diferencias en las condiciones de amplificación, con respecto a las reportadas por los autores. La amplificación de un fragmento que abarque una longitud mayor utilizando primers reportados previamente en la literatura, está en proceso de estandarización en nuestro laboratorio. Es necesario realizar nuevos estudios encaminados en la caracterización de este gen en estos hemoparásitos, teniendo en cuenta que (1) se han identificado que en otros parásitos

Apicomplexa, como Plasmodium, Theileria y Babesia, el gen 18S rRNA no se agrupa en tándem, y posee menos copias que lo reportado para otras secuencias, en comparación con como en Toxoplasma gondii (Dalrymple 1990; Guay et al. 1992; Kibe et al. 1994;

McCutchan et al. 1995); (2) para géneros como Plasmodium, se ha reportado que de acuerdo a la fase del ciclo de vida los 18S rRNA varían, de acuerdo a las necesidades del parásito (Corredor and Enea 1994); y (3) en otros organismos como bacterias, se ha demostrado que la estructura de los rRNA es más conservada que la secuencia a nivel evolutivo (Fox and Woese 1975; Gutell et al. 1994).

Así mismo, la utilización de información morfológica y molecular permite (1) identificar de especies cripticas; (2) generar nueva información, que contribuya al esclarecimiento de las relaciones filogenéticas de estos parásitos, (3) esclarecer la clasificación taxonómica de las especies identificadas mediante microscopia, y (4) contribuir al conocimiento de la diversidad real de hemoparásitos; en el caso de nuestro estudio, las secuencias obtenidas para Haemogregarinas y Haemocystidium sp., son diferentes que los reportados en Brasil y Perú (Pineda Catalan et al., 2013Úngari et al. 2018). 136 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

La caracterización morfológica y molecular de los hemoparásitos identificados en este estudio, contribuyó a ampliar el rango de hospederos y geográfico, para los tres géneros identificados, en Haemogregarinas y Haemocystidium este es el primer reporte en

Colombia y la especie P. vogli, mientras que, en el caso de Plasmodium, el rango de hospedero se amplió solamente para la especie Plasmodium kentropyxi. Nuevos estudios donde se evalúen más especies, además de un número de individuos mayor, para algunas de las especies muestreados en este estudio, deben ser desarrollados, con el fin de contribuir a la identificación de rango de hospederos, y especificidad de la infección.

A pesar del uso de taxonomía integrativa para la descripción de especies, en la actualidad la clasificación taxonómica de Hemosporidios en herpetos es un reto siendo común la reclasificación de especies entre géneros; uno de los ejemplos más representativos de esta situación son los géneros Haemocystidium y Haemoproteus, autores como Martinele et al.,(2016) y Javanbakht et al., (2015), clasifican a los hemosporidios identificados en tortugas terrestres como parte del género Haemoproteus, sin embargo, nuestros resultados concuerdan con los obtenidos por Maia et al., (2016) y Pineda-Catalán et al.,

(2013), los cuales apoyan la clasificación de los Haemosporidios que infectan reptiles en los que hay ausencia de merogonia eritrocitaria, como parte del género Haemocystidium, así como, su clasificación en dos subgéneros Haemocystidium y Simondia.

Finalmente, en cuanto a los efectos que los hemoparásitos pueden generar en estos hospederos, se ha reportado que en hospederos naturales no generan efectos negativos

(Brown et al. 2006); no obstante, dentro de los efectos que se han asociado a la infección por hemoparásitos, se encuentra la alteración de la morfología de la célula hospedera, generando hipertrofia, desplazamiento y deformación del núcleo de la célula; la infección con Plasmodium, se ha asociado a reducción de valores de hematocrito, hemoglobina, Capítulo 4 137 aumento de eritrocitos inmaduros (Telford 2009), además de alteraciones a nivel hormonal e inhibición reproductiva de las hembras de Sceloporus occidentalis infectadas por

Plasmodium mexicanum (Dunlap and Schall 1995); en serpientes se ha reportado que parasitemias bajas de Hepatozoon no generan efectos negativos (Caudell et al. 2002), sin embargo, en hospederos no naturales puede ser patógenos (Wozniak and Telford 1991).

Así mismo, dentro de las nuevas perspectivas para esta línea de investigación que se hicieron evidentes en este trabajo, se encuentran (1) debe investigarse, caracterizarse e identificarse los vectores asociados a la trasmisión de los hemoparásitos; (2) se debe investigar los efectos que puedan generar la infección por estos organismos en las especies hospederas en el marco de la medicina de la conservación; (3) debe aumentarse el muestro de reptiles con el fin de identificar la diversidad de hemoparásitos en el territorio, además de contribuir al conocimiento de rangos de hospederos y geográfico.

4.1 Referencia Ayala SC (1975) Malaria and hemogregarines from lizards of the Western Caribbean Islands of San Andrés and Providencia. Rev Inst Med Trop Sao Paulo 17:218–224

Ayala SC (1978) Checklist, Host Index, and Annotated Bibliography of Plasmodium from Reptiles. The Journal of Protozoology 25:87–100. doi: 10.1111/j.1550- 7408.1978.tb03874.x

Ayala S.C, D’Alessandro A, Mackenzie R., Angel D (1973) Hemoparasite Infections in 830 Wild Animals from the Eastern Llanos of Colombia. The Journal of Parasitology 59:52–59. doi: 10.2307/3278571

Ayala S.C, Spain JL (1976) A population of Plasmodium colombiense sp. n. in the iguanid lizard, Anolis Auratus. J Parasitol 62:177–189

Batalla L. L, Casas A. E, Julca R, et al (2015) Presencia de Hemoparásitos en Tortugas Motelo (Chelonoides denticulata) (Linnaeus, 1766) Comercializadas en el Mercado de Belén, Iquitos, Perú. Revista de Investigaciones Veterinarias del Perú 26:489. doi: 10.15381/rivep.v26i3.11168

Brown GP, Shilton CM, Shine R (2006) Do parasites matter? Assessing the fitness consequences of haemogregarine infection in snakes. Can J Zool 84:668–676. doi: 10.1139/z06-044 138 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Caudell JN, Whittier J, Conovera MR (2002) The effects of haemogregarine-like parasites on brown tree snakes (Boiga irregularis) and slatey-grey snakes (Stegonotus cucullatus) in Queensland, Australia. In: International biodeterioration & biodegradation. Elsevier, pp 113–119

Corredor V, Enea V (1994) The small ribosomal subunit RNA isoforms in Plasmodium cynomolgi. Genetics 136:857–865

Dunlap KD, Schall JJ (1995) Hormonal alterations and reproductive inhibition in male fence lizards (Sceloporus occidentalis) infected with the malarial parasite Plasmodium mexicanum . Physiological Zoology 608–621

Fox GE, Woese CR (1975) The architecture of 5S rRNA and its relation to function. J Mol Evol 6:61–76 Capítulo 4 139

Goes VC, Brito ES, Valadao RM, et al (2018) Haemogregarine (Apicomplexa: Adeleorina) infection in Vanderhaege’s toad-headed turtle, Mesoclemmys vanderhaegei (Chelidae), from a Brazilian Neotropical savanna region. FOLIA PARASIT 65:. doi: 10.14411/fp.2018.012

Guay J-M, Huot A, Gagnon S, et al (1992) Physical and genetic mapping of cloned ribosomal DNA from Toxoplasma gondii: primary and secondary structure of the 5S gene. Gene 114:165–171. doi: 10.1016/0378-1119(92)90570-F

Gutell RR, Larsen N, Woese CR (1994) Lessons from an evolving rRNA: 16S and 23S rRNA structures from a comparative perspective. Microbiol Rev 58:10–26

Gutierrez G (2016) PARÁSITOS SANGUÍNEOS EN ALGUNOS REPTILES DE LA LOCALIDAD DE YONDÓ, ANTIOQUIA, COLOMBIA. Universidad Nacional de Colombia

Javanbakht H, Kvičerová J, Dvořáková N, et al (2015) Phylogeny, Diversity, Distribution, and Host Specificity of Haemoproteus spp. (Apicomplexa: Haemosporida: Haemoproteidae) of Palaearctic Tortoises. Journal of Eukaryotic Microbiology 62:670–678. doi: 10.1111/jeu.12227

Martinele I, Tostes R, Castro R, D’Agosto M (2016) Prevalence of Haemoproteus spp. (Apicomplexa: Haemoproteidae) in tortoises in Brazil and its molecular phylogeny. Parasitology Research 115:249–254. doi: 10.1007/s00436-015-4741-3

McCutchan TF, Li J, McConkey GA, et al (1995) The cytoplasmic ribosomal RNAs of Plasmodium spp. Parasitol Today (Regul Ed) 11:134–138

Medina-Rangel GF, Méndez-Galeano MA, Calderón Espinosa ML (2019) Herpetofauna of San José del Guaviare, Guaviare, Colombia. Biota Colombiana 20:75–90. doi: 10.21068/c2019.v20n01a05

Moreno A, González L, Barreto D (2015) CARACTERIZACIÓN MORFOLÓGICA DE HEMOPARÁSITOS PRESENTES EN ALGUNOS REPTILES Y ANFIBIOS DE GUAVIARE, COLOMBIA. Universidad Colegio Mayor de Cundinamarca

Pedrozo Prieto R, Vetter Hiebert R, Quintana Ruiz Díaz A, et al (2016) HEMATOLOGIC VALUES IN CAPTIVITY TERRESTRIAL TURTLES (Chelonoidis chilensis) IN ASUNCION AND SAN LORENZO CITIES, PARAGUAY. Compend cienc vet 6:28– 35. doi: 10.18004/compend.cienc.vet.2016.06.02.28-35 140 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Perkins SL, Keller AK (2001) Phylogeny of Nuclear Small Subunit rRNA genes of Hemogregarines Amplified With Specific Primers. Journal of Parasitology 87:870– 876. doi: 10.1645/0022-3395(2001)087[0870:PONSSR]2.0.CO;2

Perkins SL, Martin JM (1999) Conserved polymerase chain reaction primers fail in diagnosis of parasitic infections. J Parasitol 85:982–984

Telford SR (1984) Haemoparasites of Reptiles. In: Hoff GL, Frye FL, Jacobson ER (eds) Diseases of Amphibians and Reptiles. Springer US, Boston, MA, pp 385–517

Telford SRJr (2009) Hemoparasites of the Reptilia: color atlas and text. Taylor & Francis, Boca Raton

Wozniak EJ, Telford SR (1991) The fate of Hepatozoon species naturally infecting florida black racers and watersnakes in potential mosquito and soft tick vectors, and histological evidence of pathogenicity in unnatural host species. International Journal for Parasitology 21:511–516. doi: 10.1016/0020-7519(91)90055-C Capítulo 4 141

Wozniak EJ, Telford SRJ, McLaughlin (1994) Employment of the polymerase chain reaction in the molecular differentiation of reptilian haemogregarines and its applications to preventative zoological medicine. Journal of Zoo and Wildlife Medicine 25:538–547

5. Conclusiones y recomendaciones

5.1 Conclusiones Los artículos desarrollados en el marco de esta tesis aportan nueva información sobre la diversidad de hemoparásitos del phylum Apicomplexa en herpetos de diversas localidades de Colombia, y su relación con géneros descritos en aves y mamíferos. Así mismo, contribuyen al esclarecimiento de las relaciones filogenéticas de géneros como

Haemocystidium; la descripción de nuevas especies con base en características morfológicas e información genética, y la obtención de información genética para especies descritas previamente con base en caracteres morfológicos.

Este estudio muestra los Llanos Orientales de Colombia como un área potencial para la investigación de hemoparásitos en herpetos, teniendo en cuenta la diversidad de fauna presente en esta zona, y la conservación de las características del territorio, podría contribuir a una biodiversidad de hemoparásitos. Primero, con base en las características morfológicas y la información molecular obtenida se reportá la presencia de

Haemocystidium sp. infectando Podocnemis vogli; segundo, se amplió el rango geográfico para Plasmodium kentropyxi y P. carmelinoi, previamente reportado en Brasil; tercero, nuevas relaciones parásito-hospedero fueron encontradas para Haemogregarina sp. infectando a P. vogli. y P. unifilis.

En relación con los hospederos, este estudio aporta información relevante para especies que poseen distribuciones restringidas o que son endémicas en nuestro país, por lo cual el poder identificar los hemoparásitos que los infectan, aportan información de base para la evaluación de efectos en el hospedero o la relación parásito-vector-hospedero vertebrado. El desarrollo de programas de conservación de las especies analizadas en el estudio, además de las amenazas de origen antropogénico deben evaluar la presencia de 143 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE Título de la tesis o trabajo de investigación DIFERENTES DEPARTAMENTOS DE COLOMBIA. microorganismos en estos hospederos, teniendo en cuenta que podrían llegar a presentar patologías asociadas a la infección, o convertirse en un pool zoonótico, en el cual organismos que no hayan estado en contacto con el parásito, se vean afectadas.

Con respecto a la distribución de los taxones descritos, a excepción de P. kentropyxi y P. carmelinoi, reportado previamente en Brasil, los demás hemoparásitos identificados son diferentes a los reportados anteriormente; en el caso de Haemogregarina sp. y

Haemocystidium sp., nuestros resultados es necesario complementar los resultados obtenidos con nuevos estudios.

El uso de la taxonomía integrativa resulta muy útil al momento de describir especies y, como en nuestro caso, la identificación de posibles especies cripticas; descartar infecciones abortivas, a través del análisis de extendidos sanguíneos, así como la identificación de coinfecciones. El uso de secuencias del citocromo b, en el caso de los

Haemosporidios, permitió caracterizar especies descritas previamente por morfología, al aportar información que corrobora la validez taxonómica del género Haemocystidium sp.

En el caso de la utilización de secuencias del 18S rRNA, permitió junto con las características morfológicas, agrupar los linajes obtenidos como parte del género

Haemogregarina, sin embargo, es necesario la utilización de nuevos marcadores moleculares de copia única que aporten nueva información que contribuyan a la resolución filogenética del suborden Adeleorina.

Conclusiones 144

5.2 Recomendaciones

(1) En el caso de los hemoparásitos pertenecientes al suborden Adeleorina, deben ser

explorados nuevos marcadores moleculares nucleares, mitocondriales, o del

apicoplasto, para mejorar resolución filogenética, y diferenciación intra e inter

especie.

(2) La realización de nuevos estudios, encaminados a identificar los vectores

asociados a la trasmisión de estos hemoparásitos, el ciclo de vida de estos

hemoparásitos y los efectos a la infección que puedan presentarse, contribuirán al

entendimiento de la relación parásito-hospedero, además, de ser información de

base para el desarrollo de programas de manejo y conservación de estas especies.

(3) La determinación de la parasitemias en reptiles debe ser estandarizada, como en

el caso de las aves; de acuerdo a la revisión de litertura realizadas, se identificó

que, de acuerdo al autor, este parámetro, ha sido evaluado en 10.000 eritrocitos o

en 2000 eritrocitos.

A. Anexo: Divulgación

Presentación Oral

González-Camacho Leydy Paola; Gutiérrez-Liberato, Germán Alfredo; Joménez, Andrés D; Vargas-Ramirez, Mario; Rodríguez-Fandiño, Oscar A; Matta-Camacho, Nubia Estela Matta. (2018) HEMOPARÁSITOS EN PODOCNEMIS VOGLI (MULLER, 1935) EN LA ORINOQUÍA COLOMBIANA; Z1-III SIMPOSIO DEL CONOCIMIENTO DE LA BIODIVERSIDAD DE PARÁSITOS EN VIDA SILVESTRE, V CONGRESO COLOMBIANO DE ZOOLOGIA, Bogotá, Colombia.

147 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE Título de la tesis o trabajo de investigación DIFERENTES DEPARTAMENTOS DE COLOMBIA.

G.A. Gutierrez, L. P Gonzalez, A. Giraldo, M. Calderon, M. Vargas, O. Rodríguez-Fandiño, N. E. Matta. Blood parasites in Neotropical herpetofauna: a huge potential of research” 4th International Conference on Malaria and Related Haemosporidian Parasites of Wildlife, at Beijing Normal University in Beijing, China, from November 1-6, 2018

Anexo A. Nombrar el anexo A de acuerdo con su contenido 148

Posters

González Leydy Paola; Moreno Andrea Marisol; Pacheco María Andreina; Escalante Ananias; González Angie Daniela; Espinosa Calderón Martha; Matta Nubia Estela; Plasmodium spp: LA DIVERSIDAD DESCONOCIDA EN LA HERPETOFAUNA DE GUAVIARE COLOMBIA. FIRST INTERNATIONAL CONGRESS OF SCIENCE, TECHONOLOGY AND INNOVATION OF THE AMERICAS. Abstract book (2017), p. 39.

Libro.

El galápago sabanero: estudios en búsqueda de su conservación/ Fundación Universitaria del Tropico Americano – Unitropico: PAREX Resources – Yopal: Dirección de investigación – Unitropico 2019 176p.:iI. Color ISBN: 4-0-52049-958-978. Capítulo. Los hemoparásitos del galápago sabanero. (Podocnemis vogli)

EL GALÁPAGO SABANERO Estudios en búsqueda de su conservación

6 7 FUNDACIÓN UNIVERSITARIA INTERNACIONAL DEL TRÓPICO AMERICANO – UNITROPICO

El galápago sabanero: estudios en búsqueda de su conservación / Fundación Universitaria Internacional del Trópico Americano – Unitrópico; PAREX Resources – Yopal: Dirección de Investigación - Unitrópico, 2019 176p.:il. Color

ISBN: 4-0-52049-958-978

Contenido: Prólogo: “Un llamado de la sábana” / Celso Román – Galápago sabanero / Oscar Andrés Rodríguez Fandiño – RNSC La Esperanza, conservación de recursos de diversas escalas / Jorge Iván Rondón Zabala Sabanas Inundables / Jorge Iván Rondón Zabala y Gilberto Augusto Cortés Millán – depresión de nido y neonatos protegidos de Podocnemis vogli (sananas inundables del Casanare del Casanare) / Gustavo Andrés López Martínez et al. – Biometría y folidosis comparada en jóvenes de tres especies del género Podocnemis (Podocnemis expansa, P. unifilis y P. vogli) de los Orientales de Colombia / Rafael Antelo Albertos – Los hemoparásitos del galápago sabanero / Leydy Paola González et al. – Taxonomía y sistemática del galápago sabanero / Gustavo López et al. – Genética de Podocnemis vogli: estado actual del conocimiento y perspectivas / Juan David Silva Galvis y Martha Lucía Ortiz Moreno – Anatomía macroscóspica y microscópica de los sistemas digestivo y respiratorio del galápago sabanero… Hernández-Henao W.J et al. -- Momentos de conservación / Jorge Iván Rondon Zabala – Bibliografías

1.TORTUGAS - 2. PODOCNEMIS – TIPOLOGÍA – 3. PODOCNEMIS – BUENAS PRÁCTICAS DE CONSERVACIÓN - I. FUNDACIÓN UNIVERSITARIA INTERNACIONAL DEL TRÓPICO AMERICANO – UNITROPICO – II. PAREX RESOURCES – III. RODRÍGUEZ FANDIÑO, Oscar Andrés – IV. CORTÉS MILLAN, Augusto Gilberto – V. RODRÍGUEZ PULIDO, José Ariel - VI. HERNÁNDEZ HENAO, William Javier – VII. MATTA CAMACHO, Nubia E – VIII. NIETO VERA, Mónica Tatiana – IX. VARGAS RAMÍREZ, Mario – X. LÓPEZ MARTÍNEZ, Gustavo Andrés – XI. JIMÉNEZ, Andrés David – XII. ANTELO ALBERTO, Rafael

CDD: 597.92 / U58

8 9 EL GALÁPAGO UN LLAMADO SABANERO DESDE LA SABANA Estudios en búsqueda de su conservación. Celso Román CONTENIDO

5 Un llamado desde la sabana. Soy el galápago sabanero, a quien los científicos bautizaron con el nombre de Podocnemis vogli, que 8 Galápago sabanero. significa caminante de Vogl, en honor del padre Cornelio Vogl, un misionero benedictino quien vivió en Venezuela y fue investigador de la Naturaleza en la Orinoquia a principios del siglo XX. 14 RNSC La esperanza y conservación de recursos en diversas escalas. Somos tortugas de cuello largo, que ante el peligro metemos la cabeza a un lado del caparazón; las 23 Las sabanas inundables, el pulso hembras medimos menos de 30 centímetros y los machos adultos son todavía más pequeños, ya de la Orinoquia que no alcanzan los 20 cm.

36 Depredación de nidos y neonatos Tal vez el padre Cornelio alcanzó a ver los numerosos grupos que formábamos en los playones protegidos de podocnemis cuando empezaba la época de anidación al llegar el verano, entre los meses de noviembre y enero. vogli (sabanas inundables del Éramos tantas, que parecíamos verdaderas invasiones descritas en 1898 por el escritor Julio Verne Casanare). en su libro El soberbio Orinoco. 46 Biometría y folidosis comparada en jóvenes de tres especies del género Pero mucho antes de la llegada de los europeos las tortugas éramos parte de la alimentación de los Podocnemis (P. expansa, P. unifilis y pueblos indígenas. Ellos recorrían las sabanas y acampaban en los territorios donde excavábamos P. vogli) de los Llanos Orientales de nuestros nidos para aprovechar nuestra carne y nuestros huevos, que extraían en grandes cantida- Colombia. des.

66 Los hemoparásitos del galápago Las cosas empezaron a cambiar cuando llegaron los europeos y con ellos la colonización desafora- sabanero. (Podocnemis vogli), da. Hace 200 años el científico Alexander von Humbolt ya daba la alarma sobre el saqueo a que eran (Testudines: Podocnemididae). sometidas las nidadas, con el fin de producir aceite extraído de los huevos, el cual comercializaban y 83 Taxonomía y sistemática del exportaban como combustible para lámparas. galápago sabanero Con la colonización llegó también la religión con la prohibición de consumir carnes rojas en la época 90 Genética de (Podocnemis vogli): de cuaresma, aumentando el consumo de nuestra carne, considerada blanca. Vinieron también los Estado actual del conocimiento y ganados y los caballos que pisoteaban los nidos, de manera que todas esas situaciones contribuye- perspectivas. ron a disminuir nuestras poblaciones de ser centenares de miles a apenas algunos cientos a media- 106 Anatomía macroscópica y dos del año 1900. microscópica de los sistemas digestivo y respiratorio del Del mismo modo, mi casa ha cambiado. Habito en las sabanas inundables del Orinoco entre Colom- galápago sabanero (Podocnemis bia y Venezuela, en terrenos que se inundan cuando las sabanas palpitan a ritmos de inviernos y vogli) Muller, 1935. (Testudines: veranos en los departamentos de Meta, Casanare, Arauca, Vichada y Guaviare. Prefiero para vivir las Podocnemididae). lagunas, jagüeyes, esteros, remansos y pequeños charcos, caños de aguas tranquilas, e incluso los morichales.

10 11 Mis hermanas y yo aprovechamos el sol de la mañana para calentarnos sobre los troncos flotantes para luego buscar nuestro alimento de pastos y plantas acuáticas, peces, insectos, ranas y cangrejos, AUTORES sin desperdiciar algo de carroña. Al llegar la noche nos refugiamos bajo el agua, medio enterradas en el lodo. Rafael Antelo Albertos Bióloga, MSC, PHD Cuando sentimos el llamado del amor al empezar el verano, salimos del agua al atardecer y buscamos Biólogo PHD Profesora Universidad de los Llanos lugares donde el sol dé con fuerza en el día. Allí excavamos los nidos en suelos duros y arcillosos, Director Científico-Fundación Palmarito. Óscar Andrés Rodríguez Fandiño que ablandamos expulsando líquido por la cloaca, para depositar hasta 15 huevos a medida que Jorge Anthony Astwood Romero Biólogo PHD. avanza la noche. Biólogo Director de Investigación, Fundación Las nidadas quedan enterradas para que el padre Sol las incube un proceso que dura de tres a Coordinador Grupo de Estudio BIOHERP Universitaria-Unitrópico. cuatro meses, pero permanecen expuestas a muchos peligros, desde reptiles como el mato de agua Universidad de Los Llanos. José Ariel Rodríguez Pulido o lobo pollero, aves como el cari-care y el carraco, mamíferos como los zorros, el rabipelado, hasta el Gilberto Augusto Cortés Millán Biol. MSC, PHD más terrible de todos: ¡El ser humano! Biólogo M.S.C Grupo de investigación en genética y repro-

Docente, Universidad Nacional Abierta y a ducción animal (GIRGA), Universidad de los Pero afortunadamente un grupo de científicos amigos míos escribió este libro para mostrarle al mundo el gran esfuerzo que están haciendo para que nosotras, las galápagas sabaneras no vayamos Distancia, CEAD Yopal. Llanos. a desaparecer para siempre. Leydy Paola González. Jorge Iván Rondón Zabala Bact. Estudiante MSC Microbiología Estudiante Biología Ambiental El 14 de enero de 2014 en el Parque Ecotemático Wisirare, ubicado en Orocué, Casanare, nos Universidad Nacional de Colombia. Fundación Universitaria-Unitrópico. estudiaron, midieron, pesaron y compararon con mis parientes la charapa y la terecay, que vivían allí William Javier Hernández Henao. Juan David Silva Galvis en cautividad. (Q.E.P.D.) MVZ Estudiante de biología Por otra parte, en la Reserva Natural de la Sociedad Civil del Hato La Esperanza, en la sabana Coordinador Grupo de Estudio en Reptiles Universidad de los Llanos. inundable de Paz de Ariporo, también en Casanare, lograron proteger con malla metálica 102 nidos Federico Medem Universidad de Los Llanos. Mario Vargas Ramírez en tres épocas de postura, entre los años 2016 a 2018, para que una nueva generación pudiera vivir Andrés David Jiménez. Biólogo PHD. Grupo Biodiversidad y Conserva- protegida de los depredadores grandes que no atravesaban la malla, pero se colaron algunas Estudiante de Biología ción Genética. Instituto de Genética. Universi- hormigas y moscas que cobraron algunas pocas vidas de tortuguitas recién nacidas. Pero la mayoría Universidad Nacional de Colombia. dad Nacional de Colombia. volvieron a los humedales, de manos de los niños que desde entonces serán sus protectores. Gustavo Andrés López Martínez Mónica Tatiana Nieto Vera Este libro es un llamado desde la sabana donde cada día la modernización del campo pone en peligro Estudiante Biología Ambiental Bióloga. Joven investigador, Grupo de inves- los cuerpos de agua donde habitamos. Tal vez nunca alcancemos a ser de nuevo los centenares Fundación Universitaria-Unitrópico. tigación en genética y reproducción animal de miles de tortugas que en época de anidación vieron los indígenas, los conquistadores europeos, Nubia E. Matta Camacho (GIRGA). Humboldt o el padre Cornelio Vogl, el misionero que nos dio su nombre, pero gracias a los estudios Bact. M.S.C - PhD. Profesora Titular consignados en estos capítulos, los lectores podrán aprender a conocernos y es bien sabido que solo Universidad Nacional de Colombia. podemos amar aquello que conocemos. Fotografía Eduardo Martínez Parales Edwin Javier Amado Mejía En manos de ustedes, los seres humanos, queda la opción de cuidar la vida de este tesoro que somos RNSC La Esperanza. Fotógrafo los galápagos sabaneros. Helena Moya Arévalo María Camila Morales López Lic Biología MSc, Entomología. Profesional en medios audio visuales. Escrito por Celso Román, a nombre de las tortugas Podocnemis vogli. Martha Lucía Ortiz-Moreno

12 13 RESUMEN

La tortuga galápaga (Podocnemis vogli) hace parte de las 28 tortugas continentales que habitan en Colombia. Esta tortuga se distribuye en la Orinoquia colombiana y venezolana, y en la actualidad los estudios realizados en su población son limitados, ninguno de ellos ha analizado la diversidad de parásitos en esta especie. Los parásitos sanguíneos, son organismos biodiversos que se encuentran distribuidos a nivel mundial, infectando a gran variedad de mamíferos, aves y reptiles. En las tortugas, los protozoarios pertenecientes al phyla Apicomplexa más comúnmente encontrados son aquellos del orden Adeleina (géneros Haemogregarina y Hemolivia); también como parásitos pertenecientes al orden Haemosporida (géneros Plasmodium, Haemoproteus y Haemocystidium). Este es el primer estudio realizado en esta especie, que buscó caracterizar la biodiversidad de parásitos sanguíneos presentes en la especie Podocnemis vogli en dos localidades de la Orinoquia Colombiana. Dicho conocimiento es importante para comprender aspectos de la biología de esta especie y las relaciones tróficas con su entorno natural. Con el fin de LOS HEMOPARÁSITOS conocer dicha diversidad, realizamos diferentes muestreos en los departamentos de Casanare (Municipio de Caño Chiquito: Paz de Ariporo) y Vichada (Municipio de Puerto DEL GALÁPAGO SABANERO Carreño); para cada tortuga de rio se tomó una muestra de sangre, con la cual se reali- (PODOCNEMIS VOGLI) (TESTUDINES: PODOCNEMIDIDAE). zaron extendidos sanguíneos en láminas y se almacenó muestra sanguínea para poste- riores análisis moleculares (genéticos) del parásito. Por microscopía, se encontró que la totalidad de los 103 individuos analizados estaban infectados con parásitos Haemogregari- Leydy P. González \ Nubia Estela Matta Camacho nas; de dichos parásitos se identificaron tres formas morfológicas o morfotipos. También Andrés David Jiménez \ Oscar Andrés Rodríguez Fandiño se reporta por primera vez en Colombia y por primera vez en esta especie de tortuga el parásito Haemocytidium sp.; este parásito se presentó en co-infección con Haemogregari- nas en tres individuos provenientes de Puerto Carreño-Vichada. Estos resultados demuestran el gran potencial de investigación en tortugas, y abre nuevos retos para caracterizar e identificar como: ¿cuáles son las consecuencias de tales infecciones en las tortugas?; ¿Afecta esta infección el periodo de vida o la capacidad de postura? ¿Quién está trans- mitiendo dicho parásito?; ¿Cuál es la identidad a especie de cada morfotipo encontrado? ¿Es el Haemocystidium encontrado en Podocnemis vogli, similar al reportado en Podoc- nemis unifilis y P. expansa de Perú? ¿Es posible que otros reptiles que cohabitan con dicha tortuga tengan el mismo parásito?

72 73 GENERALIDADES DE LOS PARÁSITOS SANGUÍNEOS PUNTOS CENTRALES DEL CAPÍTULO DEL GALÁPAGO SABANERO

Los parásitos sanguíneos del género Haemogregarina son muy frecuentes Prácticamente toda especie animal presenta algún grado de parasitismo [1]. Esta relación en el galápago sabanero que habita la Orinoquia Colombiana. El estudio de interespecífica se compone de una notable multiplicidad de adaptaciones y extraordinaria hemoparásitos en tortugas de vida silvestre aporta información relevante diversidad biológica, donde los parásitos y sus hospederos se interrelacionan de una forma para ser considerados en los programas de conservación. muy íntima en sus procesos evolutivos e interactúan constantemente a través de sus historias de vida. Resulta obvio que estudiar la totalidad de los parásitos de un hospedero desborda las capacidades de los investigadores, por lo que el presente esfuerzo se centra en un grupo particular (parásitos sanguíneos) ampliamente distribuido (prácticamente en todos los hábitats a nivel global); compuesto por una gran diversidad de especies y con estrategias de transmisión muy diversas, desde transmisión directa hasta complejos procesos tróficos [2], datos con los que se puede obtener información importante del hospedero y su relación con el entorno.

Aunque dentro de la extensa diversidad de hemoparásitos se han registrado virus, bacterias, protozoarios y metazoarios, debido a su eficiencia de transmisión y capacidad de adaptación, los protozoarios del phyla Apicomplexa son las especies más registradas en la literatura. Dichos parásitos, han sido reportados en anfibios, reptiles, peces, aves y mamíferos [3]. Dentro de estos hemoprotozoarios dos importantes grupos se han encontrado infectando frecuentemente tortugas: las Haemogregarinas (Danilewsky 1885) [4-7], y los hemosporidios (Danilewsy 1885) [4, 8, 9]. Estos parásitos poseen un ciclo de vida complejo heteroxeno (involucrando más de un hospedero en su ciclo de vida), con un hospedero intermediario vertebrado (en este caso la tortuga) y un hospedero invertebrado que cumple funciones de vector; que para las tortugas de río al parecer son las sanguijuelas, en el caso de la transmisión de las Haemogregarinas [3, 10] y pudieran estar involucrados garrapatas y dípteros para los hemosporidos [3]. En el vertebrado, los parásitos desarrollan una parte de su ciclo de vida en los tejidos (en órganos como el hígado) y otra en sangre periférica (formas que se transmiten al vector). El vector, se infecta al ingerir formas parasitarias de la tortuga (llamadas gametocitos o gamontes) y dentro de él ocurren

74 75 usualmente fases de desarrollo sexual que terminan con formas infectivas en sus partes METODOLOGÍA bucales, y una vez el vector vuelve a alimentarse, ocurre la transmisión del parásito a otra tortuga. COMÓ SE PROCESÓ A PODOCNEMIS VOGLI

De los parásitos mencionados anteriormente, en el orden Adeleina, se encuentran los géneros Haemogregarina y Hemolivia, los cuales son los parásitos sanguíneos más comúnmente registrados en las especies de tortugas alrededor del globo [11-13], pero en especial, el género Haemogregarina es el más común en las tortugas de río neotropicales [5, 7]. Por otro lado, dentro de los hemosporidos también conocidos como parásitos causantes de malaria se han reportado los géneros Plasmodium, Haemoproteus y Haemocystidium infectando tortugas [4, 8, 9, 14].

Cabe resaltar, que aunque el estudio de los parásitos en especies y poblaciones en vida silvestre se remonta a varias décadas atrás [15, 16], solo hasta hace muy pocos años se empezó a explorar la diversidad parasítica en especies del género Podocnemis, en especial en el amazonas brasileño y recientemente en Perú, donde habitan otras especies del género de gran importancia en conservación como es el caso de Podocnemis expansa y P. unifilis [6, 7, 9, 17].

Dado que los parásitos sanguineos han demostrado tener un impacto sobre la calidad de vida de sus hospederos pues entre otros aspectos pueden disminuir su capacidad reproductiva, de movilidad o actividad, incluso anemia [12, 18-20]. A partir de lo anterior, consideramos que la exploración de los parásitos de las tortugas de río es un aspecto fundamental para comprender aspectos de la biología de las tortugas en su entorno natural. Más aún en esta especie el galápago sabanero, donde para conocimiento de los autores no existen registros previos de El estudio se llevó a cabo en áreas de sabanas de inundación de este tipo de estudios. los departamentos de Casanare (Municipio de Caño Chiquito: Paz de Ariporo) y Vichada (Municipio de Puerto Carreño: Finca Flores Rojas) en la Orinoquía colombiana. Estas regiones se caracterizan por poseer regímenes hidroclimáticos y de suelos que hacen que las condiciones en campo y de paisaje sean notablemente distintas entre el periodo de lluvias (comprendido entre los meses de marzo a octubre) y la temporada seca (noviembre a febrero), lo que regula diferentes aspectos del galápago sabanero, como su

76 77 ciclo reproductivo o su distribución, y a su relación parásito-hospedero de la Universidad En la salida de campo realizada en octubre a la Reserva la Esperanza, se logró colectar 11 vez la facilidad de recolecta. Por lo anterior, Nacional de Colombia-Sede Bogotá (GERPH) y sanguijuelas que estaban infestando algunas de las tortugas capturadas. El proceso de identi- la recolección de los individuos se realizó un microscopio Nikon DM1000 de la Fundación ficación y detección de parásitos aún está en proceso. principalmente en los periodos de transición Universitaria Unitrópico. Las láminas positivas, de baja precipitación a altas lluvias, debido a fueron fotografiadas con una cámara Olympus Para la recolecta y movilización del material biológico, el estudio está amparado por el que el paisaje “pulsatil” del perfil de agua D27 acoplada al microscopio y procesadas permiso Marco de recolección de especímenes silvestres de la diversidad biológica para fines superficial presenta una focalización del por medio del software CellSens Standart. La no comerciales otorgado a la Universidad Nacional de Colombia, por parte de la autoridad recurso hídrico y por tanto la mayor determinación morfológica se realizó como nacional de licencias ambientales (ANLA), resolución 0255 del 12 de marzo de 2014 y resolución concentración de tortugas y facilidad para sugiere Telford [12], Cook et al. [22], Lainson R. modificatoria 1482 del 20 de noviembre de 2015. Todos los procedimientos a realizar en este su recolección. Los ejemplares de P. vogli [8], Pineda-Catalan et al. [9], Maia J. P. [23] y proyecto han sido avalados por el comité de ética de la Fundación Universitaria Unitrópico y el se capturaron principalmente mediante Úngari et al. [7]. comité de ética de la Universidad Nacional de Colombia. recolección manual o el uso de artes de pesca (e.g. trasmallo) para su posterior manejo en campo. Los ejemplares capturados fueron medidos (morfometría estándar) [21] procesados por medio de toma de muestra sanguínea por venupunción de la vena caudal, marcados con una muesca en el margen del escudo [21] y finalmente liberados.

Con la sangre se realizaron extendidos sanguíneos, para luego ser fijados en metanol absoluto y coloreados con Giemsa para su observación micros- cópica. De igual forma se preservaron muestras sanguíneas en EDTA o etanol, para su posterior análisis molecular (análisis en proceso). La observación de los extendidos sanguíneos se realizó en un microscopio Olympus CX41 (Olympus Cor- poration, Tokyo, Japan), en el laboratorio

78 79 LOS PARÁSITOS SANGUÍNEOS ENCONTRADOS EN EL GALÁPAGO SABANERO

Como se mencionó previamente, a nivel mundial el hemoparásito más común en tortugas pertenece al género Haemogregarina, en este estudio el 100% de las tortugas analizadas estuvieron infectadas (90 provinieron de la RNSC La Esperanza-Casanare y 13 de Puerto Carreño-Vichada), y tres de ellas Figura 1. Morfotipo 1 de Hae- capturadas en Puerto Carreño, presentaron co-infección Haemocystidium y Haemogregarina. mogregarinas encontrados en las galápagas estudiadas. (A-D) Gamontes maduros. Bar 10 µm

HAEMOGREGARINAS: El mofotipo 2, se caracteriza porque su núcleo se ubica en la parte A pesar que las Haemogregarinas, son los parásitos más comunes en tortugas, la mayoría central del cuerpo del parásito, de los estudios a nivel mundial, se refieren a morfotipos (al igual que la información aquí siendo mucho más pequeño que el consignada), teniendo en cuenta que, para poder definir especies, es necesario conocer las mencionado previamente (Figura diferentes características de su ciclo de vida, incluyendo: i. Las características morfológicas 2). El citoplasma es agranular y se ubica en la célula en la misma e información de linajes moleculares de las formas parasitarias presentes en sangre (en el posición (polar) del morfotipo 1, hospedero vertebrado) y ii. las formas de desarrollo en el vector (hospedero invertebrado). generando también el desplazamiento del núcleo de la célula Cada uno de los morfotipos encontrados se diferenció con base en las características morfológicas hospedera (eritrocito). de los gamontes (que son las formas sexuales del parásito, presentes en sangre de la tortuga), como la ubicación y forma de su núcleo y la apariencia del citoplasma, así como las diferencias en su tamaño (morfometría-datos no mostrados). Dadas las diferencias en las formas de los Figura 2. Morfotipo 2. de Haemogregarinas parásitos acá llamadas morfotipos y una vez puedan ser asociadas a sus características genéticas encontrados en las galápagas estudiadas. (A-D) Gamontes maduros. Bar 10 µm. podrían tratarse de especies de parásitos diferentes. Finalmente, el morfotipo 3 (el menos prevalente) (Figura 3) es el más grande El morfotipo 1 el más prevalente (Figura 1.), se caracteriza porque su núcleo se encuentra de los tres. Sus características morfoló- ubicado en la periférica del parásito, ocupando casi la tercera parte de este (Figura 1). En sus gicas, difieren notablemente de los dos morfotipos anteriores: el citoplasma es formas inmaduras, se caracteriza por dar la apariencia de media luna (el citoplasma, no ocupa granular, en la parte anterior del parásito la totalidad del cuerpo del parásito), a medida que el parásito crece, el citoplasma va ocupando el citoplasma es rosado, en todas las ese espacio, no obstante, la posición del núcleo siempre es lateral. En cuanto a su ubicación formas observadas. Los gamontes ob- en la célula hospedera (eritrocito), el parásito se ubica hacia los polos de la misma y genera el servados, causan aumento del tamaño y desplazamiento del núcleo en los glóbulos desplazamiento de su núcleo (generalmente ubicado en el centro de la célula). rojos y en algunos, el parásito ha expulsa- do totalmente el núcleo (Figura 3C).

80 81 ANÁLISIS DE LOS RESULTADOS Y POTENCIALIDADES A FUTURO

La infección generalizada por parte del parásito, representada en una prevalencia del 100%, es un resultado muy poco común en las poblaciones de vida libre; sin embargo, la relación Hae- mogregarina – tortuga presenta unas prevalencias notablemente altas a nivel global, reportándose en diversas regiones geográficas y para diversas especies de tortugas valores superiores al 70% [29,28,11], igual para poblaciones neotropicales y de tortugas podocnemídidas [7, 9, 17].

Aunque explicar una relación tan amplia y generalizada entre estos parásitos y su hospedero

Figura. 3. Morfotipo 3. de Haemogregarinas encontrados en las galápagas estudiadas. es compleja, un primer elemento a tener en cuenta es la potencial capacidad de infectar a su (A-D) Gamontes maduros. Bar 10 µm hospedero por periodos particularmente largos. Este fenómeno se ha registrado en otros parásitos de géneros cercanos como Hemolivia y Hepatozoon, parasitando tortugas y cocodrilos, respectivamente [30, 31]. Es importante resaltar que el periodo de vida de los glóbulos rojos HAEMOCYSTIDIUM de tortugas es en promedio de 600 – 800 días [32], esto pudiera explicar porque la prevalencia tan alta, pues el recambio de eritrocitos es bajo y una vez infectado, el investigador tiene un largo periodo de tiempo donde pudiera detectarlo en sangre periférica. A lo largo de la historia la clasificación de los hemoparásitos pertenecientes a este género ha sido confusa, pues ha sido asignado a otros géneros como Plasmodium o Haemoproteus [24- De igual forma, la exposición del hospedero (tortuga) a un vector (mosquito o sanguijuela) 26]. De acuerdo a los análisis genéticos realizados hasta la fecha [9, 27], este parásito infecta especificamente a tortugas y es genéticamente diferente a Plasmodium o Haemoproteus. potencial es importante, para ello es necesario resaltar los pulsos de inundación y sequía, en En la sangre periférica de las 3 tortugas galapas infectadas, se encontraron diversos estadios especial de los cuerpos de agua lénticos que durante la temporada seca lleva a la concentración del desarrollo de este parásito desde formas iniciales (Trofozoitos) hasta las formas sexuales de los individuos de galápago sabanero [33, 34], potencialmente acercando individuos infectados que ingiere e infectan al vector (mosquito o sanguijuela) llamados gametocitos (Fig 4. A). con no infectados. Aunque se desconoce el vector para la relación Haemogregarina – Podocnemis, los únicos vectores implicados en otros estudios en el mundo son las sanguijuelas [11]. De suceder la transmisión de los parásitos en el galápago sabanero de la misma manera, este

Fig 4. Haemocystidium spp. en tortugas galapas provenientes de Puerto Carreño – Vichada. (A) forma inicial-Trofozoito, (B-C) Gametocito inmaduro (célula sexual) (D) gametocito femenino. Bar 10 µm.

82 83 vector vería optimizado el acceso a recurso alimentario con el establecimiento de la AGRADECIMIENTOS temporada seca. En este sentido, hubié- El desarrollo del presente estudio no podría ser sin el apoyo económico de Colciencias semos esperado encontrar sanguijuelas mediante la financiación del proyecto FP44842-18-2018. La Universidad Nacional de Colombia adheridas a los galápagos en las capturas y el apoyo de insfraestructura del Laboratorio Relación-Parásito-Hospedero. Al convenio de marzo y abril, sin embargo, en dichas interinstitucional entre Unitrópico y Parex Resources “Huella Galápaga”, por el apoyo económico recibido en campo en Casanare y a la Fundación Universitaria – Unitrópico. El apoyo invaluable recolectas no se observaron estos ectopa- en la fase de campo de la familia Martínez-Parales y Rafael Gutiérrez D. rásitos, mientras que en diversos ejemplares recolectados en Casanare en el mes de octubre (transición de lluvias a seca), fueron BIBLIOGRAFÍA observadas sanguijuelas en rostro y extre- 1. Poulin, R. & S. Morand, The diversity of parasites. a new species of Haemogregarina Danilewsky 1885 midades de los galápagos sabaneros (datos The quarterly review of biology, 2000. 75(3): p. 277-293. (Adeleina: Haemogregarinaidae) in free-living and no mostrados). 2. Votýpka, J., D. Modrý, M. Oborník, J. Šlapeta, captive yellow-spotted river turtles Podocnemis unifilis & J. Lukeš, Apicomplexa, in Handbook of the Protists, (Testudines: Podocnemididae) from Brazil. Parasitology J. Archibald, A. Simpson, and C. Slamovits, Editors. 2016, Research, 2018. 117(5): p. 1535-1548. A la fecha no existe un estudio específico Springer International Publishing: Netherlands. p. 1-58. 8. Lainson, R., Atlas of protozoan parasites of the Amazo- que analice el impacto de Haemogregarinas 3. O’Donoghue, P., Haemoprotozoa: Making biological nian fauna of Brazil. Vol. 1. Haemosporida of reptiles, vol. 1. . sobre esta especie de tortuga, algunos autores sense of molecular phylogenies. International Journal for 2012, Brasil: Ananindeua. Instituto Evandro Chagas. 9. Pineda-Catalan, O., S. Perkins, M. Peirce, reportan baja virulencia de dichos parásitos Parasitology: Parasites and Wildlife, 2017. 6(3): p. 241-256. R. Engstrand, C. Garcia-Davila, M. Pine- 4. Lainson, R. & R. Naiff, Haemoproteus (Apicomple- [7]. Sin embargo, es deseable analizar el do-Vasquez, & A. Aguirre, Revision of Hemopro- xa: Haemoproteidae) of Tortoises and Turtles. Proceedings impacto de estos parásitos en sus tejidos, teid Genera and Description and Redescription of Two of the Royal Society of London B: Biological Sciences, Species of Chelonian Hemoproteid Parasites. Journal y realizar estudios de seguimiento a largo 1998. 265: p. 941-949. of Parasitology, 2013. 99(6): p. 1089-1098. plazo que permitan determinar el impacto 5. Rossow, J., S. Hernandez, S. Sumner, B. 10. Davies, A. & M. Johnston, The biology of some Altman, C. Crider, M. Gammage, K. Segal, sobre su calidad de vida y éxito reproductivo. intraerythrocytic parasites of fishes, amphibia and reptiles. & M. Yabsley, Haemogregarine infections of three Advances in Parasitology, 2000. 45: p. 1-107. species of aquatic freshwater turtles from two sites Este es el primer estudio de Hemoparásitos in Costa Rica. International Journal for Parasitology: 11. Siddall, M. & S. Desser, Prevalence and intensity en el galapago sabanero en Colombia, con Parasites and Wildlife, 2013. 2: p. 131-135. of Haemogregarina balli (Apicomplexa: Adeleina: Hae- mogregarinidae) in three turtle species from Ontario, with resultados sorprendentes desde el punto 6. De Oliveira, J., M. André, J. Alves Júnior, A. Lustosa, & K. Werther, Molecular detection observations on intraerythrocytic development. Canadian de vista de prevalencia y la biodiversidad de of hemogregarines and haemosporidians in Brazilian Journal of Zoology, 1992. 70(1): p. 123-128. parásitos encontrados. Estudios de identifi- free-living testudines. International Journal for Parasi- 12. Telford, S., Hemoparasites of the Reptilia: color atlas cación de la información genética de dichos tology: Parasites and Wildlife, 2018. 7(1): p. 75-84. and text. 2009, Boca Raton, Florida.: CRC Press. 13. Alhaboubi, A., D. Pollard, & P. Holman, Molecu- parásitos y su potencial vector deben ser 7. Úngari, L., A. Santos, L. O’Dwyer, M. da Silva, N. de Melo Fava, G. Paiva, & M. Cury, lar and morphological characterization of a haemogregari- adelantados. Haemogregarina podocnemis sp. nov.: description of ne in the alligator snapping turtle, Macrochelys temminckii

84 85 (Testudines: Chelydridae). Parasitology Research, 2017. in reptiles in Faculdade de Ciências. 2015, Universida- 116(1): p. 207-215. de do Porto: Porto, Portugal. 14. Picelli, A., A. Carvalho, L. de Viana, & A. Mal- 24. Wenyon, C., The pigmented parasites of cold-blooded vasio, Prevalence and parasitemia of Haemogregarina sp. animals, with some notes on a Plasmodium of the Trinidad in Podocnemis expansa (Testudines: Podocnemididae) from iguana. Journal of Tropical Medicine and Hygiene, 1915. 18: the Brazilian Amazon. Revista Brasileira de Parasitologia p. 133-140. Veterinária, 2015. 24(2): p. 191-197. 25. Wenyon, C., Protozoology : a manual for medi- 15. Ayala, S., A. D’Alessandro, R. Mackenzie, & D. cal men, veterinarians and zoologists. 1926, New York: Angel, Hemoparasite Infections in 830 Wild Animals from William Wood & Company. the Eastern Llanos of Colombia. The Journal of Parasitology, 26. Mackerras, M., The haematozoa of Australian repti- 1973. 59(1): p. 52-59. les. Australian Journal of Zoology, 1926. 9: p. 61-122. 16. Ayala, S., Malaria and hemogregarines from 27. Maia, J., D. Harris, S. Carranza, & E. Goméz- lizards of the Western Caribbean Islands of San Andrés Díaz, Assessing the diversity, host-specificity and infection and Providencia. Revista Do Instituto De Medicina Tropi- patterns of apicomplexan parasites in reptiles from Oman, cal De São Paulo, 1975. 17(4): p. 218-224. Arabia. Parasitology, 2016. 143(13): p. 1730-1747. 17. Soares, P., E. de Brito, F. Paiva, D. Pavan, & L. 28. Davis, A. & S. Sterrett, Prevalence of haemo- Viana, Haemogregarina spp. in a wild population from Po- gregarine parasites in three freshwater turtle species docnemis unifilis Troschel, 1848 in the Brazilian Amazonia. in a population in northeast Georgia, USA. International Parasitology research, 2014. 113(12): p. 4499-4503. journal of zoological Research, 2011. 7(2): p. 156-163. 18. Dunlap, K. & J. Schall, Hormonal alterations 29. Jakes, K., P. O’Donoghue, M. Munro, & and reproductive inhibition in male fence lizards (Sce- R. Adlard, Hemoprotozoa of freshwater turtles in loporus occidentalis) infected with the malarial parasite Queensland. Journal of Wildlife Diseases, 2001. 37(1): p. Plasmodium mexicanum Physiological Zoology, 1995. 12-19. 68(4): p. 608–621. 30. Široký, P., M. Kamler, & D. Modrý, Long-term 19. Bouma, M., C. Smallridge, C. Bull, & J. occurrence of Hemolivia cf. mauritanica (Apicomplexa: Komdeur, Susceptibility to infection by a haemo- Adeleina: Haemogregarinidae) in captive Testudo marginata gregarine parasite and the impact of infection in the (Reptilia: Testudinidae): evidence for cyclic merogony? Jour- Australian sleepy lizard Tiliqua rugosa. Parasitology nal of Parasitology, 2004. 90(6): p. 1391-1393. Research, 2007. 100(5): p. 949-954. 31. Viana, L., F. Paiva, M. Coutinho, & R. Lou- 20. Oppliger, A. & J. Clobert, Reduced tail regene- renço-de-Oliveira, Hepatozoon caimani (Apicomplexa: ration in the common lizard, Lacerta vivipara, parasitized Hepatozoidae) in wild caiman, Caiman yacare, from the Pan- by blood parasites. Functional Ecology, 1997. 11(5): p. tanal region. Journal of Parasitology, 2010. 96(1): p. 83-88. 652-655. 32. Berlin, N., T. Waldmann, & S. Weissman, 21. Rueda-Almonacid, J.V., L. Carr, R. Mitter- Life span of red blood cell. Physiological Reviews, 1959. meier, J. Rodríguez-Mahecha, R. Mast, R. Vogt, 39(3): p. 577-616. A. Rhodin, J. de la Ossa-Velásquez, J. Rueda, 33. Alarcon-Pardo, H., Contribución al conocimiento de & C. Mittermeier, Las tortugas y los cocodrilianos de la morfología, ecologia, comportamiento y distribución geo- los países andinos del trópico. Serie de guías tropicales gráfica de Podocnemis vogli, Testudinata (Pelomedusidae). TAXONOMÍA de campo Nº 6. 2007, Bogotá: Conservación Internacio- Revista de la Academia Colombiana de Ciencias Exactas nal. Editorial Panamericana. 538. Físicas y Naturales, 1969. 13: p. 303-329. Y SISTEMÁTICA 22. Cook, C., N. Smit, & A. Davies, Haemoproteids 34. Ramo, C., Biología del galápago (Podocnemis vogli, (Apicomplexa: Haemoproteidae) from South African tor- Müller, 1935) en el Hato» El Frío», Llanos de Apure (Vene- DEL GALÁPAGO toises (Cryptodira: Testudinidae). Journal of Parasitology, zuela). Acta Vertebrata, 1982. 9(3): p. 1-161. 2010. 96: p. 1168–1172. SABANERO 23. Maia, J., Diversity, infection patterns and host–parasite associations of apicomplexan parasites Gustavo López, Jorge Rondón, Mario Vargas-Ramírez, Oscar Rodríguez

86 87 EL GALÁPAGO SABANERO Estudios en búsqueda de su conservación.

176 177 EL GALÁPAGO SABANERO Estudios en búsqueda de su conservación 166 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE Título de la tesis o trabajo de investigación DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Bibliografía Adl, S. M., Leander, B. S., Simpson, A. G., Archibald, J. M., Anderson, O. R., Bass, D., ... & Kolisko, M. (2007). Diversity, nomenclature, and taxonomy of protists. Systematic Biology, 56(4), 684-689.

Adl, S.M., Simpson, A.G.B., Lane, C.E., Lukeš, J., Bass, D., Bowser, S.S., Brown, M.W., Burki, F., Dunthorn, M., Hampl, V., Heiss, A., Hoppenrath, M., Lara, E., le Gall, L., Lynn, D.H., McManus, H., Mitchell, E.A.D., Mozley-Stanridge, S.E., Parfrey, L.W., Pawlowski, J., Rueckert, S., Shadwick, L., Schoch, C.L., Smirnov, A., Spiegel, F.W., 2012. The Revised Classification of Eukaryotes. Journal of Eukaryotic Microbiology 59, 429–514. https://doi.org/10.1111/j.1550-7408.2012.00644.x

Adl, S.M., Bass, D., Lane, C.E., Lukeš, J., Schoch, C.L., Smirnov, A., Agatha, S., Berney, C., Brown, M.W., Burki, F., Cárdenas, P., Čepička, I., Chistyakova, L., del Campo, J., Dunthorn, M., Edvardsen, B., Eglit, Y., Guillou, L., Hampl, V., Heiss, A.A., Hoppenrath, M., James, T.Y., Karpov, S., Kim, E., Kolisko, M., Kudryavtsev, A., Lahr, D.J.G., Lara, E., Le Gall, L., Lynn, D.H., Mann, D.G., Massana i Molera, R., Mitchell, E.A.D., Morrow, C., Park, J.S., Pawlowski, J.W., Powell, M.J., Richter, D.J., Rueckert, S., Shadwick, L., Shimano, S., Spiegel, F.W., Torruella i Cortes, G., Youssef, N., Zlatogursky, V., Zhang, Q., 2019. Revisions to the Classification, Nomenclature, and Diversity of Eukaryotes. Journal of Eukaryotic Microbiology. https://doi.org/10.1111/jeu.12691

Allen KE, Yabsley MJ, Johnson EM, et al (2011) Novel Hepatozoon in VertebratesFrom the Southern United States. Journal of Parasitology 97:648–653. doi: 10.1645/GE- 2672.1

Alley M, Hale K, Cash W, et al. (2010) Concurrent avian malaria and avipox virus infection in translocated South Island saddlebacks (Philesturnus carunculatus carunculatus). N Z Vet J 58:218–223. doi: 10.1080/00480169.2010.68868

Altschul, S.F., Madden, T.L., Schäffer, A.A., Zhang, J., Zhang, Z., Miller, W., Lipman, D.J., 1997. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 25, 3389–3402.

Andrade C MG (2011) Estado del conocimiento de la biodiversidad en Colombia y sus amenazas. Consideraciones para fortalecer la interacción ambiente-política. Rev Acad Colomb Cienc 35:491–507

Ayala S.C (1970) Plasmodium mexicanum in California: natural history and development in Phlebotomine sandflies (Diptera: Psychodidae). J Parasitol 56:13 Bibliografía 167

Ayala SC (1975) Malaria and hemogregarines from lizards of the Western Caribbean Islands of San Andrés and Providencia. Rev Inst Med Trop Sao Paulo 17:218–224

Ayala S.C, D’Alessandro A, Mackenzie R., Angel D (1973) Hemoparasite Infections in 830 Wild Animals from the Eastern Llanos of Colombia. The Journal of Parasitology 59:52–59. doi: 10.2307/3278571

Ayala S.C, Spain JL (1976) A population of Plasmodium colombiense sp. n. in the iguanid lizard, Anolis Auratus. J Parasitol 62:177–189

Ayala SC (1978) Checklist, Host Index, and Annotated Bibliography of Plasmodium from Reptiles. J Protozool 25:87–100. doi: 10.1111/j.1550-7408.1978.tb03874.x

Ayala SC, Hertz PE (1981) Malaria infection in Anolis lizards on Martinique, Lesser Antilles. Rev Inst Med Trop Sao Paulo 23:12–17

Barta JR (1991) The Dactylosomatidae. In: Advances in Parasitology. Elsevier, pp 1–37

Bensch, S., Stjernman, M., Hasselquist, D., Ostman, O., Hansson, B., Westerdahl, H., Pinheiro, R., 2000. Host specificity in avian blood parasites: a estudy of Plasmodium and Haemoproteus mitochondrial DNA amplifield from birds. Proc Biol Sci 267, 1583– 1589.

Benson, D.A., Karsch-Mizrachi, I., Clark, K., Lipman, D.J., Ostell, J., Sayers, E.W., 2012. GenBank. Nucleic Acids Research 40, D48–D53. https://doi.org/10.1093/nar/gkr1202

Bonorris JS, Ball GH (1955) Schellackia occidentalis n.sp., a Blood-inhabiting Coccidian Found in Lizards in Southern California. J Protozool 2:31–34. doi: 10.1111/j.1550- 7408.1955.tb02393.x

Bouet, J., 1909. Sur deux he mocytö zaires pigmente s̈ des reptiles. C.R. Se ä nc. Soc. Biol. 66, 43–45. 168 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE Título de la tesis o trabajo de investigación DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Boundenga, L., Makanga, B., Ollomo, B., Gilabert, A., Rougeron, V., Mve-Ondo, B., Arnathau, C., Durand, P., Moukodoum, N.D., Okouga, A.-P., Delicat-Loembet, L., Yacka-Mouele, L., Rahola, N., Leroy, E., Ba, C.T., Renaud, F., Prugnolle, F., Paupy, C., 2016. Haemosporidian Parasites of Antelopes and Other Vertebrates from Gabon, Central Africa. PLOS ONE 11, e0148958. https://doi.org/10.1371/journal.pone.0148958

Boundenga, L., Perkins, S.L., Ollomo, B., Rougeron, V., Leroy, E.M., Renaud, F., Prugnolle, F., 2017. Haemosporidian Parasites of Reptiles and Birds from Gabon, Central Africa. Journal of Parasitology 103, 330–337. https://doi.org/10.1645/16-118

Brown GP, Shilton CM, Shine R (2006) Do parasites matter? Assessing the fitness consequences of haemogregarine infection in snakes. Can J Zool 84:668–676. doi: 10.1139/z06-044

Castaño-Mora, 2002. Libro rojo de reptiles de Colombia. La serie Libros rojos de especies amenazadas.

Castellani, A., Willey, A., 1904. Observations on the haematozoa of vertebrates in Ceylon. Spolia Zeylonic 2, 78–92.

Craig TM, Smallwood JE, Knauer KW, McGrath JP (1978) Hepatozoon canis infection in dogs: clinical, radiographic, and hematologic findings. J Am Vet Med Assoc 173:967– 972

Cohen SC, Justo MCN, Kohn A (2013) South American Monogenoidea parasites of fishes, amphibians and reptiles. South American Monogenoidea parasites of fishes, amphibians and reptiles Bibliografía 169

Cook, C.A., Smit, N.J., Davies, A.J., 2010. Hemoproteids (Apicomplexa: Haemoproteidae) from South African Tortoises (Cryptodira: Testudinidae). Journal of Parasitology 96, 1168–1172. https://doi.org/10.1645/GE-2527.1

Corredor V, Enea V (1994) The small ribosomal subunit RNA isoforms in Plasmodium cynomolgi. Genetics 136:857–865

Darriba D, Taboada GL, Doallo R, Posada D (2012) jModelTest 2: more models, new heuristics and parallel computing. Nat Methods 9:772–772. doi: 10.1038/nmeth.2109

Davies AJ, Johnston MRL (2000) The biology of some intraerythrocytic parasites of fishes, amphibia and reptiles. In: Parasitology B-A in (ed). Academic Press, pp 1–107 DeGiusti, D.L., Sterling, C.R., Dobrzechowski, D., 1973. Transmission of the Chelonian Haemoproteid Haemoproteus metchnikovi by a Tabanid Fly Chrysops callidus. Nature 242, 50–51. https://doi.org/10.1038/242050a0

Davis AK, Sterrett SC (2011) Prevalence of Haemogregarine Parasites in Three Freshwater Turtle Species in a Population in Northeast Georgia, USA. International J of Zoological Research 7:156–163. doi: 10.3923/ijzr.2011.156.163 170 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE Título de la tesis o trabajo de investigación DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Daszak P (2000) Emerging Infectious Diseases of Wildlife-- Threats to Biodiversity and Human Health. Science 287:443–449. doi: 10.1126/science.287.5452.443

DeGiusti, D.L., Sterling, C.R., Dobrzechowski, D., 1973. Transmission of the Chelonian Haemoproteid Haemoproteus metchnikovi by a Tabanid Fly Chrysops callidus. Nature 242, 50–51. https://doi.org/10.1038/242050a0 de Oliveira JP, André MR, Alves Júnior JR, et al (2018) Molecular detection of hemogregarines and haemosporidians in Brazilian free-living testudines. International Journal for Parasitology: Parasites and Wildlife 7:75–84. doi: 10.1016/j.ijppaw.2018.01.008

Drummond AJ, Ho SYW, Phillips MJ, Rambaut A (2006) Relaxed Phylogenetics and Dating with Confidence. PLoS Biol 4:e88 . doi: 10.1371/journal.pbio.0040088

Edgar RC (2004) MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Res 32:1792–1797 doi: 10.1093/nar/gkh340

Escalante, A.A., Freeland, D.E., Collins, W.E., Lal, A.A., 1998. The evolution of primate malaria parasites based on the gene encoding cytochrome b from the linear mitochondrial genome. Proc. Natl. Acad. Sci. U.S.A. 95, 8124–8129.

Escobar S. F (2000) Diversidad de coleopteros coprofagos (Scarabaeidae: Scarabaeinae) en un mosaico de habitats en la reserva natural Nukak, Guaviare, Colombia. Acta Zool Mex 103–121

Falk BG, Glor RE, Perkins SL (2015) Clonal reproduction shapes evolution in the lizard malaria parasite Plasmodium floridense. Evolution 69:1584–1596. doi: 10.1111/evo.12683

Fernandes BMM, Kohn A (2014) South American trematodes parasites of amphibians and reptiles. South American trematodes parasites of amphibians and reptiles

Fox GE, Woese CR (1975) The architecture of 5S rRNA and its relation to function. J Mol Evol 6:61–76

Gaffney, E.S., Meylan, P.A., Wood, R.C., Simons, E., De Almeida Campos, D., 2011. Evolution of the side-necked turtles: the family Podocnemididae. Bulletin of the American Museum of natural History.

Garnham PCC (1966) Malaria Parasites and other Haemosporidia. Blackwell Scientific Publications, Oxford, U. K

Garcés-Restrepo MF, Giraldo A, Carr JL, Brown LD (2013) Turtle ectoparasites from the Pacific coastal region of Colombia. Biota Neotrop 13:74–79. doi: 10.1590/S1676- 06032013000300009 172 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE Título de la tesis o trabajo de investigación DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Gutell RR, Larsen N, Woese CR (1994) Lessons from an evolving rRNA: 16S and 23S rRNA structures from a comparative perspective. Microbiol Rev 58:10–26

Gutierrez G (2016) PARÁSITOS SANGUÍNEOS EN ALGUNOS REPTILES DE LA LOCALIDAD DE YONDÓ, ANTIOQUIA, COLOMBIA. Universidad Nacional de Colombia

Hellgren, O., Križanauskiene, A., Valkiūnas, G., Bensch, S., 2007. Diversity and phylogeny of mitochondrial cytochrome b lineages from six morphospecies of avian Haemoproteus (Haemosporida: Haemoproteidae). Journal of Parasitology 93, 889– 896. https://doi.org/10.1645/GE-1051R1.1

Hellgren O, Waldeström J, Bensch S (2004) A new PCR assay for simultaneous studies of Leucocytozoon, Plasmodium, and Haemoproteus from avian blood. J Parasitol 90:797–802 Bibliografía 173

Herrera, J.R., 2008. Estudio patológico retrospectivo de mortalidad en reptiles del zoológico Jaime Duque entre el año 1991 y el 2006. Universidad de la Salle, Bogotá, D.C.

Haklová B, Majláthová V (2014) Phylogenetic relationship of Hepatozoon blood parasites found in snakes from Africa, America and Asia. Parasitology 141:389–398. doi: 10.1017/S0031182013001765

Hoffman (1999) Parasites of North American freshwater fishes. University of California Press, Berkeley, Los Angeles, London

Hoffmann M, Hilton-Taylor C, Angulo A, et al (2010) The Impact of Conservation on the Status of the World’s Vertebrates. Science 330:1503–1509. doi: 10.1126/science.1194442

Jakes, K.A., O’Donoghue, P., Munro, M., Adlard, R., 2001. Hemoprotozoa of freshwater turtles in Queensland. Journal of Wildlife Diseases 37, 12–19. https://doi.org/10.7589/0090-3558-37.1.12

Jarvi, S.I., Schultz, J.J., Atkinson, C.T., 2002. PCR diagnostics underestimate the prevalence of avian malaria (Plasmodium relictum) in experimentally-infected passerines. Journal of Parasitology 88, 153–158. https://doi.org/10.1645/0022- 3395(2002)088[0153:PDUTPO]2.0.CO;2

Javanbakht, H., Kvičerová, J., Dvořáková, N., Mikulíček, P., Sharifi, M., Kautman, M., Maršíková, A., Široký, P., 2015. Phylogeny, Diversity, Distribution, and Host Specificity of Haemoproteus spp. (Apicomplexa: Haemosporida: Haemoproteidae) of Palaearctic Tortoises. Journal of Eukaryotic Microbiology 62, 670–678. https://doi.org/10.1111/jeu.12227

Johnston, T.H., Cleland, J.B., 1909. On a new melanin- producing haematozoon from an Australian tortoise. J. Proc. R. Soc. NSW 43, 97–103.

Kibe, M. K., Nene, V., Khan, B., Allsopp, B. A., Collins, N. E., Morzaria, S. P., ... & Bishop, R. P. (1994). Evidence for two single copy units in Theileria parva ribosomal RNA genes. Molecular and biochemical parasitology, 66(2), 249-259. 174 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE Título de la tesis o trabajo de investigación DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Krasilnikov, E.N., 1965. Parazity krovy cherepakh yugo- vostochnoi Gruzii (Blood parasites of turtles of southeast Georgia). Zool. Zh. 44, 1454–1459.

Kumar, S., Stecher, G., Tamura, K., 2016. MEGA7: Molecular Evolutionary Genetics Analysis Version 7.0 for Bigger Datasets. Molecular Biology and Evolution 33, 1870– 1874. https://doi.org/10.1093/molbev/msw054

Lainson R, Landau I, Shaw JJ (1974) Observations on non-pigmented haemosporidia of Brazilian lizards, including a new species of Saurocytozoon in Mabuya mabouya (Scincidae). Parasitology 69:215–223

Lainson R, Shaw JJ, Landau I (1975) Some blood parasites of the Brazilian lizards Plica umbra and Uranoscodon superciliosa (Iguanidae). Parasitology 70:119–141. doi: 10.1017/S0031182000048927

Lainson, R., Naiff, R.D., 1998. Haemoproteus (Apicomplexa: Haemoproteidae) of Tortoises and Turtles. Proceedings: Biological Sciences 265, 941–949.

Lainson R (2012) Atlas of protozoan parasites of the amazonian fauna of Brazil. Vol. 1. Haemosporida of reptiles.

Larkin MA, Blackshields G, Brown NP, et al (2007) Clustal W and Clustal X version 2.0. Bioinformatics 23:2947–2948. doi: 10.1093/bioinformatics/btm404 Bibliografía 175

Laverna, 1905. Sur une he mamibë nouvelle de Testudo pardalis. C. R. Soc. Se anc.̈ Biol. 59, 176-178}.

Le Blancq, S. M., Khramtsov, N. V., Zamani, F., Upton, S. J., & Wu, T. W. (1997). Ribosomal RNA gene organization in Cryptosporidium parvum. Molecular and biochemical parasitology, 90(2), 463-478.

Levine ND (1970) Taxonomy of the sporozoa. J Parasitol 56:208–209

Levine, ND (1988) The protozoan phylum Apicomplex. p. 118-133. 1988. CRC Press, INC, Boca Raton, Florida

Levine, N.D., 1988. The protozoan phylum Apicomplex. p. 118-133. 1988. CRC Press, INC, Boca Raton, Florida.

Levine, N.D., 1970. Taxonomy of the sporozoa. J. Parasitol. 56, 208–209.

Lotta, I.A., Pacheco, M.A., Escalante, A.A., González, A.D., Mantilla, J.S., Moncada, L.I., Adler, P.H., Matta, N.E., 2016. Leucocytozoon Diversity and Possible Vectors in the Neotropical highlands of Colombia. Protist 167, 185–204.

Mackerras, M., 1961. The Haematozoa of Australian Reptiles. Australian Journal of Zoology 9, 61. https://doi.org/10.1071/ZO9610061

Maia JP (2015) Diversity, infection patterns and hostparasite associations of apicomplexan parasites in reptiles. (Doctoral dissertation, Ph. D. Thesis. Universidade do Porto, Porto, Portugal).

Maia, D. JamesHarris, Salvador Carranza (2016) Reconstruction of the evolutionary history of Haemosporida (Apicomplexa) based on the cyt b gene with characterization of 176 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE Título de la tesis o trabajo de investigación DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Haemocystidium in geckos (Squamata: Gekkota) from Oman. Parasitology international, 65:5–11

Mantilla, J.S., Matta, N.E., Pacheco, M.A., Escalante, A.A., González, A.D., Moncada, L.I., 2013. Identification of Plasmodium (Haemamoeba) lutzi (Lucena, 1939) from Turdus fuscater (great thrush) in Colombia. The Journal of parasitology 99, 662–668.

Matta, N.E., Pacheco, M.A., Escalante, A.A., Valkiūnas, G., Ayerbe-Quiñones, F., Acevedo-Cendales, L.D., 2014. Description and molecular characterization of Haemoproteus macrovacuolatus n. sp. (Haemosporida, Haemoproteidae), a morphologically unique blood parasite of black-bellied whistling duck (Dendrocygna autumnalis) from South America. Parasitology research 113, 2991–3000.

Martinele, I., Tostes, R., Castro, R., D’Agosto, M., 2016. Prevalence of Haemoproteus spp. (Apicomplexa: Haemoproteidae) in tortoises in Brazil and its molecular phylogeny. Parasitology Research 115, 249–254. https://doi.org/10.1007/s00436-015-4741-3

McCutchan, T. F., Li, J., McConkey, G. A., Rogers, M. J., & Waters, A. P. (1995). The cytoplasmic ribosomal RNAs of Plasmodium Spp. Parasitology Today, 11(4), 134- 138.

Medina-Rangel GF, Méndez-Galeano MA, Calderón Espinosa ML (2019) Herpetofauna of San José del Guaviare, Guaviare, Colombia. Biota Colombiana 20:75–90. doi: 10.21068/c2019.v20n01a05 Bibliografía 177

Mercereau-Puijalon, O., Barale, J. C., & Bischoff, E. (2002). Three multigene families in Plasmodium parasites: facts and questions. International journal for parasitology, 32(11), 1323-1344.

Misra, K.K., Choudhury, A., 1977. Haemoproteus trionyxii n.sp. from a river turtle, Trionyx gangeticus Cuvier. Arch. Protistenkd. 119, 395–400.

Moore, R. B, Oborník, M, Janouškovec, J, et al (2008) A photosynthetic alveolate closely related to apicomplexan parasites. Nature 451:959

Morand S, Krasnov BR, Littlewood DTJ (eds) (2015) Parasite Diversity and Diversification: Evolutionary Ecology Meets Phylogenetics. Cambridge University Press, Cambridge

Moreno A, González L, Barreto D (2015) CARACTERIZACIÓN MORFOLÓGICA DE HEMOPARÁSITOS PRESENTES EN ALGUNOS REPTILES Y ANFIBIOS DE GUAVIARE, COLOMBIA. Universidad Colegio Mayor de Cundinamarca

Moreno LA, Andradre GI, Ruíz-Contreras LF, (Eds.) (2016) Biodiversidad 2016. Estado y tendencia de la biodiversidad de Colombia.

Morrison, D.A., 2009. Evolution of the Apicomplexa: where are we now? Trends in Parasitology 25, 375–382. https://doi.org/10.1016/j.pt.2009.05.010

Muehlenbein, M.P., Pacheco, M.A., Taylor, J.E., Prall, S.P., Ambu, L., Nathan, S., Alsisto, S., Ramirez, D., Escalante, A.A., 2015. Accelerated Diversification of Nonhuman Primate Malarias in Southeast Asia: Adaptive Radiation or Geographic Speciation? Molecular Biology and Evolution 32, 422–439. https://doi.org/10.1093/molbev/msu310

Murata T, Inoue M, Tateyama S, et al (1993) Vertical transmission of Hepatozoon canis in dogs. J Vet Med Sci 55:867–868 178 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE Título de la tesis o trabajo de investigación DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Murphy J, Jowers M (2013) Treerunners, cryptic lizards of the Plica plica group (Squamata, Sauria, Tropiduridae) of northern South America. ZooKeys 355:49–77. doi: 10.3897/zookeys.355.5868

Morales-Betancourt MA, Lasso CA, Páez VP, Bock BC (2015) Libro rojo de reptiles de Colombia. Instituto de Investigación de Recursos Biológicos Alexander Von Humboldt, Colombia

Morand S, Krasnov BR, Littlewood DTJ (eds) (2015) Parasite diversity and diversification: evolutionary ecology meets phylogenetics. Cambridge University Press, Cambridge, United Kingdom

Morrison DA (2009) Evolution of the Apicomplexa: where are we now? Trends in Parasitology 25:375–382. doi: 10.1016/j.pt.2009.05.010

Oppliger A, Célérier ML, Clobert J (1996) Physiological and behaviour changes in common lizards parasitized by haemogregarines. Parasitology 113:433–438. doi: 10.1017/S003118200008149X

Orkun, Ö., Güven, E., 2013. A New Species of Haemoproteus from a Tortoise (Testudo graeca ) in Turkey, with Remarks on Molecular Phylogenetic and Morphological Analysis. Journal of Parasitology 99, 112–117. https://doi.org/10.1645/GE-3100.1 Bibliografía 179

Outlaw, D.C., Ricklefs, R.E., 2014. Species limits in avian malaria parasites (Haemosporida): how to move forward in the molecular era. Parasitology 141, 1223– 1232. https://doi.org/10.1017/S0031182014000560

Ortiz-Catedral L, Brunton D, Stidworthy MF, et al (2019) Haemoproteus minutus is highly virulent for Australasian and South American parrots. Parasites Vectors 12:40. doi: 10.1186/s13071-018-3255-0

Ortiz-Moreno ML, Rodríguez-Pulido JA (2017) Estado del conocimiento y amenazas de la tortuga sabanera (Podocnemis vogli, Podocnemididae) en Colombia. Orinoquia 21:26. doi: 10.22579/20112629.391

Pachón, D.A., 2009. Aislamiento, identificación y serotipificación de enterobacterias del género Salmonella en una población Crocodylus intermedius y testudinos mantenidos en cautiverio en la estación Biológica Roberto Franco E.B.T.R.B de la Facultad de Ciencias- Universidad Nacional de Colombia. Pontificia Universidad Javeriana, Bogotá, D.C.

Pacheco, M., Cranfield, M., Cameron, K., Escalante, A.A., 2013. Malarial parasite diversity in chimpanzees: the value of comparative approaches to ascertain the evolution of Plasmodium falciparum antigens. Malaria Journal 12, 328. https://doi.org/10.1186/1475-2875-12-328

Pacheco, M.A., Cepeda, A.S., Bernotienė, R., Lotta, I.A., Matta, N.E., Valkiūnas, G., Escalante, A.A., 2018. Primers targeting mitochondrial genes of avian haemosporidians: PCR detection and differential DNA amplification of parasites belonging to different genera. International Journal for Parasitology 48, 657–670. https://doi.org/10.1016/j.ijpara.2018.02.003

Pacheco, M.A., Matta, N.E., Valkiunas, G., Parker, P.G., Mello, B., Stanley, C.E., Lentino, M., Garcia-Amado, M.A., Cranfield, M., Kosakovsky Pond, S.L., Escalante, A.A., 2018. Mode and Rate of Evolution of Haemosporidian Mitochondrial Genomes: Timing the Radiation of Avian Parasites. Mol. Biol. Evol. 35, 383–403. https://doi.org/10.1093/molbev/msx285

Pachón D, Pulido V A, Moreno T C (2011) Aislamiento y serotipificación de Salmonella sp. en estanques con Crocodylus intermedius y testudines cautivos en Villavicencio - Colombia. Rev MVZ Córdoba 16:2564. doi: 10.21897/rmvz.1021 180 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE Título de la tesis o trabajo de investigación DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Palinauskas, V., Žiegytė, R., Ilgūnas, M., Iezhova, T.A., Bernotienė, R., Bolshakov, C., Valkiūnas, G., 2015. Description of the first cryptic avian malaria parasite, Plasmodium homocircumflexum n. sp., with experimental data on its virulence and development in avian hosts and mosquitoes. International Journal for Parasitology 45, 51–62. https://doi.org/10.1016/j.ijpara.2014.08.012

Perkins SL, Martin JM (1999) Conserved polymerase chain reaction primers fail in diagnosis of parasitic infections. J Parasitol 85:982–984

Perkins, 2000. Species concepts and malaria parasites: detecting a cryptic species of Plasmodium. Proceedings of the Royal Society of London. Series B: Biological Sciences 267, 2345–2350. https://doi.org/10.1098/rspb.2000.1290

Perkins SL (2001) Phylogeography of Caribbean lizard malaria: tracing the history of vector- borne parasites. J Evol Biol 14:34–45

Perkins SL, Schall J (2002) A molecular phylogeny of malarial parasites recovered from cytochrome b gene sequences. J Parasitol 88:972–978

Perkins SL, Austin CC (2009) Four new species of Plasmodium from New Guinea lizards: integrating morphology and molecules. J Parasitol 95:424–433

Perkins SL (2014) Malaria’s Many Mates: Past, Present, and Future of the Systematics of the Order Haemosporida. Journal of Parasitology 100:11–25. doi: 10.1645/13-362.1 Bibliografía 181

Perkins, S.L., Martinsen, E.S., Falk, B.G., 2011. Do molecules matter more than morphology? Promises and pitfalls in parasites. Parasitology 138, 1664–1674. https://doi.org/10.1017/S0031182011000679

Pittaluga, J., 1912. Ein neuer Blutparasit de afrikanischer Schildröte, Clemmys africana, Haemoproteus cajale n.sp. Centralbl. f. Bakt. Parasitkde 63, 241–243.

Plimmer, H.G., 1912. On the blood parasites found in animals in the Zoological Gardens during the four years 1908-1911. Proc. Zool. Soc. Lond. 27, 406–419.

Ricklefs RE, Fallon SM, Bermingham E (2004) Evolutionary relationships, cospeciation, and host switching in avian malaria parasites. Syst Biol 53:111–119

Rhodin, A.G.J., Iverson, J.B., Bour, R., Fritz, U., Georges, A., Shaffer, H.B., van Dijk, P.P., 2017. Turtles of the World: Annotated Checklist and Atlas of Taxonomy, Synonymy, Distribution, and Conservation Status (8th Ed.). Chelonian Research Foundation & Turtle Conservancy. https://doi.org/10.3854/crm.7.checklist.atlas.v8.2017

Ronquist, F., Huelsenbeck, J.P., 2003. MrBayes 3: Bayesian phylogenetic inference under mixed models. Bioinformatics 19, 1572–1574. https://doi.org/10.1093/bioinformatics/btg180

Rossow, J.A., Hernandez, S.M., Sumner, S.M., Altman, B.R., Crider, C.G., Gammage, M.B., Segal, K.M., Yabsley, M.J., 2013. Haemogregarine infections of three species of aquatic freshwater turtles from two sites in Costa Rica. International Journal for Parasitology: Parasites and Wildlife 2, 131–135. https://doi.org/10.1016/j.ijppaw.2013.02.003 182 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE Título de la tesis o trabajo de investigación DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Rueda-Almonacid, Carr, J. L., Mittermeier, R. A, Rodríguez-Mahecha, J. V., Mast, R. B., Vogt, R. C., Mittermeier, C. G., 2007. Las tortugas y los cocodrilianos de los países andinos del trópico. Serie de guías tropicales de campo, 6, 412–423.

Sambrook, J., Fritsch, E.F., Maniatis, T., 1989. Molecular cloning. Cold spring harbor laboratory press New York.

Schall JJ (1983) Lizard malaria: cost to vertebrate host’s reproductive success. Parasitology 87:1–6. doi: 10.1017/S0031182000052367

Schall JJ (2002) Parasite Virulence. Behav. Ecol. Parasites 283–313

Schneider CA, Rasband WS, Eliceiri KW (2012) NIH Image to ImageJ: 25 years of image analysis. Nat Methods 9:671–675. doi: 10.1038/nmeth.2089

Sehgal RNM, Hull AC, Anderson NL, et al. (2006) Evidence for cryptic speciation of Leucocytozoon spp. (Haemosporida, Leucocytozoidae) in diurnal raptors. J Parasitol 92:375–379

Siddall ME, Burreson EM (1994) The Development of a Hemogregarine of Lycodes raridens from Alaska in Its Definitive Leech Host. The Journal of Parasitology 80:569. doi: 10.2307/3283193

Simond, P.L., 1901. Contribution á l’e tudë des he matozoaires̈ endoglobulaires des reptils. Ann. Inst. Pasteur 15, 319–351.

Smith TG (1996) The genus Hepatozoon (Apicomplexa: Adeleina). J Parasitol 82:565–585

Staats, C.M., Schall, Jos.J., 1996. Distribution and Abundance of Two Malarial Parasites of the Endemic Anolis Lizard of Saba Island, Netherlands Antilles. The Journal of Parasitology 82, 409–413. https://doi.org/10.2307/3284077 184 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE Título de la tesis o trabajo de investigación DIFERENTES DEPARTAMENTOS DE COLOMBIA.

Stacy NI, Alleman AR, Sayler KA (2011) Diagnostic Hematology of Reptiles. Clin Lab Med 31:87–108 . doi: 10.1016/j.cll.2010.10.006

Tamura, K., Nei, M., 1993. Estimation of the number of nucleotide substitutions in the control region of mitochondrial DNA in humans and chimpanzees. Molecular Biology and Evolution. https://doi.org/10.1093/oxfordjournals.molbev.a040023

Telford, S.R.Jr., 1996. Two new species of Haemocystidium Castellani & Willey (Apicomplexa: Plasmodiidae) from Pakistani lizards, and the support their meronts provide for the validity of the genus. Systematic Parasitology 34, 197–214. https://doi.org/10.1007/BF00009387

Telford SR Jr. (1973) Malaria Parasites of the “Borriguerro’’Lizard, Ameiva ameiva (Sauria: Teiidae) in Panama. J Protozool 20:203–207. doi: 10.1111/j.1550- 7408.1973.tb00863.x

Telford, S.R., 1984. Haemoparasites of Reptiles, in: Hoff, G.L., Frye, F.L., Jacobson, E.R. (Eds.), Diseases of Amphibians and Reptiles. Springer US, Boston, MA, pp. 385– 517. https://doi.org/10.1007/978-1-4615-9391-1_20

Telford S.R. Jr. (1988) A Contribution to the systematics of the reptilian malaria parasites, family Plasmodiidae (Apicomplexa Haemospororina). University of Florida, Gainesville

Telford, S.R., 2005. A Haemocystidium species from the east african gecko Lygodactylus capensis grotei. Journal of Parasitology 91, 135–138. https://doi.org/10.1645/GE- 3395

Telford, S.R.J., 2007. Redescription of Haemoproteus mesnili (Apicomplexa: Plasmodiidae) and its meronts, with description of a second haemosporidian parasite of African Cobras. Journal of Parasitology 93, 673–679. https://doi.org/10.1645/GE- 3582.1

Telford, S.R.Jr., 2009. Hemoparasites of the Reptilia: color atlas and text. Taylor & Francis, Boca Raton. Bibliografía 185

Úngari, L.P., Santos, A.L.Q., O’Dwyer, L.H., da Silva, M.R.L., de Melo Fava, N.N., Paiva, G.C.M., de Melo Costa Pinto, R., Cury, M.C., 2018. Haemogregarina podocnemis sp. nov.: description of a new species of Haemogregarina Danilewsky 1885 (Adeleina: Haemogregarinaidae) in free-living and captive yellow-spotted river turtles Podocnemis unifilis (Testudines: Podocnemididae) from Brazil. Parasitology Research 117, 1535–1548. https://doi.org/10.1007/s00436-018-5817-7

Valkiūnas, G., 2005. Avian malaria parasites and other haemosporidia. CRC Press, Boca Raton.

Valkiūnas, G., Iezhova, T.A., 2018. Keys to the avian malaria parasites. Malaria Journal 17, 212. https://doi.org/10.1186/s12936-018-2359-5

Valkiūnas, G., Iezhova, T.A., Križanauskienė, A., Palinauskas, V., Sehgal, R.N.M., Bensch, S., 2008. A Comparative Analysis of Microscopy and PCR-Based Detection Methods for Blood Parasites. Journal of Parasitology 94, 1395–1401. https://doi.org/10.1645/GE-1570.1

Valkiūnas, G., Iezhova, T.A., Loiseau, C., Sehgal, R.N.M., 2009. Nested Cytochrome B Polymerase Chain Reaction Diagnostics Detect Sporozoites of Hemosporidian Parasites in Peripheral Blood of Naturally Infected Birds. Journal of Parasitology 95, 1512–1515. https://doi.org/10.1645/GE-2105.1

Valkiūnas, G., Sehgal, R.N.M., Iezhova, T.A., Hull, A.C., 2010. Identification of Leucocytozoon toddi Group (Haemosporida: Leucocytozoidae), with Remarks on the Species Taxonomy of Leucocytozoids. Journal of Parasitology 96, 170–177. https://doi.org/10.1645/GE-2109.1

Valkiūnas G, Z̆ic̆ kus T, Shapoval AP, Iezhova TA (2006) Effect of Haemoproteus belopolskyi (Haemosporida: Haemoproteidae) on Body Mass of the Blackcap Sylvia atricapilla. Journal of Parasitology 92:1123–1125. doi: 10.1645/GE-3564-RN.1 186 IDENTIFICACIÓN DE HEMOPARÁSITOS PRESENTES EN LA HERPETOFAUNA DE Título de la tesis o trabajo de investigación DIFERENTES DEPARTAMENTOS DE COLOMBIA.

van As, J., Cook, C.A., Netherlands, E.C., Smit, N.J., 2016. A new lizard malaria parasite Plasmodium intabazwe n. sp. (Apicomplexa: Haemospororida: Plasmodiidae) in the Afromontane Pseudocordylus melanotus (Sauria: Cordylidae) with a review of African saurian malaria parasites. Parasites & Vectors 9. https://doi.org/10.1186/s13071- 016-1702-3

Vargas-Ramirez, M., Castaño-Mora, O., Fritz, U., 2008. Molecular phylogeny and divergence times of ancient South American and Malagasy river turtles (Testudines: Pleurodira: Podocnemididae). Organisms Diversity & Evolution 8, 388–398. https://doi.org/10.1016/j.ode.2008.10.001 van Riper III C, van Riper SG, Goff ML, Laird M (1986) The Epizootiology and Ecological Significance of Malaria in Hawaiian Land Birds. Ecol Monogr 56:327. doi: 10.2307/1942550

Vardo-Zalik AM, Schall JJ (2008) Clonal diversity within infections and the virulence of a malaria parasite, Plasmodium mexicanum. Parasitology 135:1363–1372. doi: 10.1017/S0031182008004964

Velandia O (2019) La biodiversidad y los servicios ecosistémicos. http://www.humboldt.org.co/es/biodiversidad/que-es-la-biodiversidad. Accessed 2 Mar 2019

Vitt LJ, Colli GR (1994) Geographical ecology of a Neotropical lizard: Ameiva ameiva (Teiidae) in Brazil. Can J Zool 72:1986–2008. doi: 10.1139/z94-271

Votýpka, J, Modrý, D., Oborník, M., et al (2017) Apicomplexa. In: Handbook of the Protists. Springer International Publishing., pp 1–58

Walther, E.L., Valkiūnas, G., González, A.D., Matta, N.E., Ricklefs, R.E., Cornel, A., Sehgal, R.N.M., 2014. Description, molecular characterization, and patterns of distribution of a widespread New World avian malaria parasite (Haemosporida: Plasmodiidae), Plasmodium (Novyella) homopolare sp. nov. Parasitology Research 113, 3319–3332. https://doi.org/10.1007/s00436-014-3995-5 Bibliografía 187

Wenyon CM (1915) The pigmented parasites of cold-blooded animals, with some notes on a Plasmodium of the Trinidad iguana. Journal of Tropical Medicine and Hygiene 18:133–140

Wenyon CM (1926) Protozoology: a manual for medical men, veterinarians and zoologists. New York: W. Wood

Whiteman, N.K., Sánchez, P., Merkel, J., Klompen, H., Parker, P.G., 2006. Cryptic host specificity of an avian skin mite (Epidermoptidae) vectored by louse flies (Hippoboscidae) associated with two endemic Galapagos bird species. Journal of Parasitology 92, 1218–1228.

WHO, 2018. Malaria [WWW Document]. world health organization. URL https://www.who.int/es/news-room/fact-sheets/detail/malaria (accessed 3.26.19).