Epidemiología Y Control De La Fiebre Q (Coxiella Burnetii) En Fauna Silvestre Ibérica

Epidemiología y control de la fiebre Q (Coxiella burnetii) en fauna silvestre ibérica.

David González Barrio

Tesis doctoral

Epidemiología y control de la fiebre Q (Coxiella burnetii) en fauna silvestre ibérica.

Trabajo de investigación presentado por David González Barrio para optar al grado de Doctor por la Universidad de Castilla-La Mancha

Ciudad Real, 2015 Grupo de Sanidad y Biotecnología (SaBio) Instituto de Investigación en Recursos Cinegéticos (IREC; CSIC- UCLM-JCCM) Departamento de Ciencia y Tecnología Agroforestal y Genética Universidad de Castilla-La Mancha

Los abajo firmantes, como directores de este tesis doctoral, hacemos constar que la Tesis titulada “Epidemiología y control de la fiebre Q (Coxiella burnetii) en fauna silvestre ibérica”, y realizada por David González Barrio, reúne los requisitos necesarios para su defensa y aprobación y, por tanto, para optar al grado de doctor con mención internacional.

Vº Bº de los Directores

Dr José Francisco Ruiz Fons

Dr Isabel G. Fernández de Mera

Dr Christian Gortázar Schmidt

La realización de este trabajo ha sido posible gracias a las siguientes entidades y proyectos de investigación:

Fundación de la Universidad de Castilla-La Mancha

Cátedra Fundación Enresa

Proyecto Europeo Strategies For The Eradication Of Bovine Tuberculosis

Proyecto Europeo, Comisión Europea Harmonised Approaches In Monitoring Wildlife Population Health, And Ecology And Abundance (APHAEA)

Proyecto Europeo, Comisión Europea (VII Programa Marco) ANTIcipating the global onset of new epidemics (ANTIGONE)

Junta de Comunidades de Castilla-La Mancha Estructura de los contactos y riesgo de transmisión de enfermedades entre Ganado y ungulados silvestres

CGT – Apoyo Al Desarrollo De Nuevas Tecnologias Para El Control Sanitario de La Fauna Silvestre

Ministerio de Economía y Competitividad Centro para el Desarrollo Tecnológico Industrial (CDTI) – Incorporación de Nuevas Metodologías para la tecnificación y sostenibilidad de explotaciones bovinas extensivas y cinegáticas.

Financiación Adicional Del Contrato Ramón Y Cajal De José Francisco Ruiz Fons

Resumen.

Capítulo I: Introducción general.

Capítulo II: Epidemiología de Coxiella burnetii en fauna silvestre ibérica.

1. Estado de Coxiella burnetii en las poblaciones de ciervo rojo (Cervus elaphus)

en la península ibérica y factores de riesgo asociados.

“Host and Environmental Factors Modulate the Exposure of Free-Ranging

and Farmed Red Deer (Cervus elaphus) to Coxiella burnetii”

2. Estado de Coxiella burnetii en las poblaciones de conejo de monte

(Oryctolagus cuniculus) en la península ibérica y factores de riesgo asociados.

“European Rabbits as Reservoir for Coxiella burnetii”

3. Dinámica de la infección por Coxiella burnetii en una población endémica de

ciervo en condiciones semi-extensivas.

“Long-term dynamics of Coxiella burnetii in farmed red deer (Cervus

elaphus)”

4. Genotipos de Coxiella burnetii presentes en fauna silvestre en la península

ibérica basados en MLVA.

“Coxiella burnetii genotypes in Iberian wildlife”

5. Genotipado de Coxiella burnetii de fauna silvestre ibérica mediante PCR e

hibridación RLB y relaciones con genotipos de ganado doméstico y humanos

en España.

“Coxiella burnetii genotypes in Spanish wildlife: implications for livestock

and human health”

Capítulo III: Vías de transmisión de Coxiella burnetii en fauna silvestre ibérica.

1. Vías de excreción de Coxiella burnetii y otros patógenos relevantes en jabalí

(Sus scrofa).

“Shedding patterns of endemic Eurasian wild boar (Sus scrofa) pathogens”

2. Vías de excreción de Coxiella burnetii en ciervo rojo (Cervus elaphus) en

condiciones de producción semi-extensiva.

“Coxiella burnetii Shedding by Farmed Red Deer (Cervus elaphus)”

Capítulo IV: Estrategias de control de Coxiella burnetii: evaluación de la vacunación con vacunas inactivadas comerciales de fase I como estrategia de reducción de la prevalencia y el nivel de excreción de la bacteria en ciervo rojo (Cervus elaphus).

Capítulo V: Síntesis general y conclusiones.

Estructura de la tesis

La finalidad de este trabajo de investigación consiste en estudiar el papel de la fauna silvestre en la epidemiología de Coxiella burnetii y en evaluar posibles medidas de control

- que al igual que en especies domésticas, sobre todo rumiantes - pueden llevarse a cabo para disminuir la contaminación ambiental - fuente de infección para otras especies silvestres, domésticas y para las personas - y, así, la transmisión.

La presente Tesis Doctoral se estructura en una primera sección (Resumen) que pretende plasmar de forma resumida los aspectos metodológicos y resultados más destacables obtenidos en cada capítulo.

A continuación se presenta el estado actual del tema (Capítulo I) a modo de Introducción

General a los trabajos de la Tesis Doctoral, en la que se aborda el conocimiento general sobre C. burnetii, patógeno intracelular obligado con un amplio rango de hospedadores y con carácter zoonósico que lo convierte en la causa de brotes epidémicos de fiebre Q en personas. Esta introducción abarca aspectos generales del patógeno (etiología, estructura genética, etc) y de sus efectos en hospedadores donde la epidemiología y el resultado de la infección han sido bien caracterizados (principalmente rumiantes domésticos y ser humano), pero su intención fundamental es centrarse en el conocimiento existente sobre el papel que juega la fauna silvestre en la epidemiología de C. burnetii y en las estrategias de control potencialmente aplicables en fauna silvestre. Incluye aspectos relacionados con

C. burnetii en fauna silvestre como las particularidades de su diagnóstico, desde la detección serológica de anticuerpos frente a C. burnetii a la caracterización molecular de los genotipos circulantes en diversas especies silvestres a nivel mundial y la relación filogenética con los genotipos aislados en otras especies de hospedadores, así como el conocimiento existente sobre los factores de riesgo de mantenimiento y transmisión de

C. burnetii asociados a la fauna silvestre. También en esta introducción se abordan

7 aspectos de la transmisión, patogenia y patología conocidos en fauna silvestre, así como el conocimiento actual sobre los métodos de control de las enfermedades en la fauna silvestre.

El Capítulo II se centra en el abordaje científico llevado a cabo en esta Tesis Doctoral para mejorar el conocimiento de la epidemiología de C. burnetii en poblaciones de fauna silvestre ibérica, en concreto en el ciervo rojo (Cervus elaphus) y en el conejo de monte

(Oryctolagus cuniculus), así como para identificar algunos de los factores de riesgo que determinan la transmisión y mantenimiento de C. burnetii en un ciclo silvestre en estas especies. En este capítulo también se aborda un estudio que pretende comprender la dinámica temporal de C. burnetii en poblaciones de ciervo rojo endémicas, usando una granja ibérica de ciervo rojo como modelo. Dos estudios adicionales abordan el tipado molecular de las cepas de C. burnetii circulantes en especies silvestres ibéricas. En estos estudios se compara mediantes dos técnicas de tipado molecular diferentes (MLVA y

PCR-RLB) los genotipos que circulan en la fauna silvestre ibérica con cepas presentes en ganado doméstico y en casos clínicos humanos en España y en el resto del mundo.

En el siguiente capítulo (Capítulo III) se aborda el estudio de las vías de transmisión de

C. burnetii en diferentes especies de fauna silvestre ibérica, concretamente en el jabalí

(Sus scrofa) y en el ciervo rojo (Cervus elaphus), así como las potenciales implicaciones clínicas de la infección por C. burnetii en el ciervo.

El último de los estudios científicos que componen la presente Tesis Doctoral (Capítulo

IV) aborda el diseño y aplicación de estrategias potenciales de control de C. burnetii en ciervo usando como modelo una granja de ciervo ibérica. El control de C. burnetii en rumiantes domésticos se basa principalmente en el uso de vacunas inactivadas de bacterias en fase I con la finalidad de reducir la prevalencia de excreción y, en paralelo, los niveles de excreción para reducir la transmisión y, con ello, la incidencia. Este capítulo

8 se basa en el diseño de un programa de vacunación experimental en condiciones de campo y en la implementación de una vacuna comercial inactivada de C. burnetii en fase I

(COXEVAC, CEVA Santè Animale, Francia), así como en la posterior evaluación de la eficacia de la vacuna en inducir inmunidad humoral y en reducir la excreción de C. burnetii en secreciones vaginales, leche y heces en un periodo de tres años desde la implementación del programa vacunal.

Esta Tesis Doctoral se cierra con una Síntesis General (Capítulo V) y Conclusiones en la que se destacan los principales hitos logrados, la aplicabilidad de los resultados y las necesidades futuras de abordaje científico necesarias para mejorar el conocimiento sobre la epidemiología, la patogenia, la clínica y el control de C. burnetii en la fauna silvestre.

Esta sección incluye las conclusiones derivadas del trabajo de investigación desarrollado en los diferentes estudios que componen la Tesis Doctoral.

Introducción

1. El patógeno y sus características

Coxiella burnetii, previamente denominada como Rickettsia diaporica o Rickettsia burnetii es una bacteria que se encuadra dentro de la clase Gammaproteobacteria, orden

Legionellales, familia Coxiellaceae. Es un bacilo gram negativo, patógeno intracelular obligado, pequeño (0.2-0.4 μm ancho, 0.4-1.0 μm de largo), no capsulado y pleomórfico

(Maurin y Raoult, 1999). Coxiella burnetii presenta dos formas antigénicas relacionadas con mutaciones en la capa de lipopolisacáridos, mutaciones que le confieren importantes variaciones antigénicas denominadas “variación de fase”, hecho similar al que ocurre en algunas enterobacterias en el que se da la transición de un lipopolisacárido liso a rugoso y que se detecta mediante técnicas serológicas (Brezina, 1958; Hackstadt et al., 1985).

Estas formas son: i) la fase antigénica I, fase natural altamente infecciosa y virulenta, y que se aisla principalmente de animales, artrópodos o humanos infectados; y ii) la fase antigénica II, menos infecciosa y que sólo se obtiene tras repetidos pases de la fase I en medios de cultivo celular o huevos embrionados (Hackstadt, 1990; Maurin y Raoult,

1999). Existen también tres variantes celulares diferentes: a) variante celular grande (En inglés ‘Large Cell Variant’ - LCV); b) variable celular pequeña (SCV del inglés ‘Small cell variant’); y c) variante celular pequeña y densa (‘Small Dense Cell’ en inglés - SDC)

(McCaul, 1991; McCaul y Williams, 1981). Estas formas presentan distintas características morfológicas, antigénicas y metabólicas, y distinto grado de resistencia a agentes físico-químicos. Así LCV es la variante intracelular, más pleomórfica y metabólicamente activa, mientras que SDC y SCV son las formas extracelulares y tienen una morfología similar. SDC se puede visualizar dentro de las LCV en forma de endosporas que se liberarán al medio tras la lisis de las LCV o por fisión binaria. Las formas SCV se encuentran en el espacio periplasmático. La formación de estas variantes

10 es una estrategia de la bacteria para sobrevivir ya que expresan distintas proteínas específicas reconocidas por los anticuerpos, lo que permite a la bacteria escapar y sobrevivir dentro del endosoma. Las variantes pequeñas (SCV y SDC) están consideradas como formas de resistencia extracelular. Coxiella burnetii tiene una gran afinidad por los fagocitos mononucleares y por lo tanto en la fase aguda de la enfermedad (infección sistémica) podemos encontrar a la bacteria en órganos como el bazo, pulmón, hígado y médula ósea, aparte de en la sangre (Maurin y Raoult, 1999). Sin embargo, el principal

órgano de replicación de C. burnetii en animales no gestantes podría ser el bazo (Zhang et al., 2005). En animales gestantes, la bacteria muestra preferencia por los tejidos del aparato reproductivo, sobre todo en tejido que se está desarrollando y multiplicando, como las células trofoblásticas de la membrana corioalantoidea de los rumiantes, siendo también evidente la multiplicación de C. burnetii en los cotiledones de la placenta

(Sánchez et al., 2006).

Coxiella burnetii se caracteriza principalmente por su gran resistencia medioambiental debida en gran parte a la capacidad para diferenciarse en variantes celulares pequeñas que son estables en el medio ambiente. Esta forma es la fagocitada por los macrófagos durante las primeras fases de la infección. Esta variantes (endospora) presentan también una alta resistencia a agentes físicos y químicos (Babudieri, 1959). Son resistentes a la desecación, a altas temperaturas, al choque osmótico, a la luz ultravioleta y a diferentes desinfectantes como el hipoclorito al 0,5%, lysol al 5% y formol al 5% durante 24 horas a 24ºC, que no eliminan por completo a la bacteria (McCaul y Williams, 1981; Ransom y Huebner, 1951;

Scott y Williams, 1990). Otros compuestos como el etanol al 70% y el cloroformo al 5% aplicado durante 30 minutos sí son capaces de inactivar a la bacteria completamente

(Scott y Williams, 1990). La eficacia de estos desinfectantes químicos se puede ver afectada por el contenido de materia orgánica presente en el medio, procedente de los

11 tejidos, fluidos de los partos y de las heces, que podría neutralizar la acción germicida de estos productos. Coxiella burnetii puede mantenerse viable e infectiva durante 4 meses en el suelo a temperatura ambiente, en lana durante 9 meses, en agua hasta 36 meses y en heces de garrapatas puede sobrevivir hasta casi 2 años (Pascual-Velasco, 1996). También resiste las bajas temperaturas, más de dos años a -20ºC. Coxiella burnetii también ha sido detectada en diferentes productos de origen animal como huevos, mayonesa, productos lácteos - como mantequilla o queso fresco - y carne fresca (Tatsumi et al., 2006). En queso permanece hasta 42 días viable, mientras en carne fresca permanece viable hasta un mes a 4ºC. También puede permanecer viable en la ropa en condiciones de alta humedad, bajas temperaturas y sin exposición directa al sol (EFSA, 2010). Esta característica de alta resistencia explica su amplia distribución y presencia en el medio, donde un ambiente ventoso puede crear condiciones favorables para su transmisión, y su capacidad para infectar animales y humanos tras grandes periodos después de haber sido excretada

(Maurin y Raoult, 1999; Tissot-Dupont et al., 2004; Arricau-Bouvery et al., 2005). Por su alta infectividad y su transmisión por medio de aerosoles (Brooke et al., 2013; Brooke et al., 2015), y su resistencia a los cambios ambientales extremos, esta bacteria ha sido clasificada en la categoría B de armas biológicas (Madariaga et al., 2003) considerándose como potencial arma en bioterrorismo.

2. Reseña histórica

Coxiella burnetii es el agente causal de la fiebre Q, enfermedad zoonósica altamente infecciosa compartida entre animales y humanos (Maurin y Raoult, 1999). En humanos esta enfermedad se detectó en 1935 en trabajadores de un matadero después de un brote febril en Brisbane, Queensland, Australia, (Derrick, 1937); los análisis laboratoriales resultaron ser negativos a todos los patógenos conocidos hasta el momento (Babudieri,

1959). El patógeno fue investigado por E. H. Derrick, director del Laboratorio de

Microbiología y Patología del Departamento de Salud de Queensland, inoculando sangre de los trabajadores enfermos en cobayas y produciéndose en estas un cuadro febril. De estos animales se volvió a inocular sangre en otros animales produciéndose de nuevo un cuadro febril, lo que suponía la implicación de un agente infeccioso. De este modo la enfermedad que provocaba este patógeno desconocido se le denominó “Q fever” (la “Q” es la abreviatura de “query”, que en inglés quiere decir interrogación) (Babudieri, 1959).

Las investigaciones continuaron hasta que M. Burnet consiguió aislar el patógeno de animales de laboratorio; estos patógenos se identificaron como organismos similares a rickettsias, denominándolo Rickettsia burnetii (Reimer, 1993). Al mismo tiempo en

Estados Unidos, H. R. Cox y su grupo investigaba la fiebre de las montañas rocosas. Estos habían recolectado garrapatas en la localidad de Nine Mile (Montana), y se suponía que habían hallado el agente causante de la fiebre de las montañas rocosas (Derrick et al.,

1939, Smith et al., 1940). Al contrario de lo que pensaban, aislaron otro patógeno con similares características a las rickettsias, al que denominaron Rickettsia diaporica, haciendo referencia a la propiedad de estos microorganismos de pasar a través de los filtros. A la enfermedad producida por este patógeno la denominaron “fiebre de Nine

Mile”. En sucesivas investigaciones H.R. Cox enfermó con un cuadro febril igual al detectado en los trabajadores de Brisbane, permitiendo relacionar en ese momento este cuadro con la enfermedad descrita en Australia (Babudieri, 1959). La inoculación de la sangre de Cox en cobayas desarrolló la enfermedad en estos y de sus bazos se pudo aislar el agente Nine Mile. En 1948, C. B. Philip propone que Rickettsia burnetii sea considerada como una especie única de un género distinto, y propone el nombre de

Coxiella burnetii para el agente causal de la fiebre Q (Marrie, 1990), en honor a Cox y

Burnet. El cuadro clínico característico de la fiebre Q humana descrito por Derrick en su primer trabajo, fue detallado y ampliado posteriormente por el mismo autor (Derrick,

1973). Desde la identificación de C. burnetii como agente causante de la fiebre Q, esta bacteria ha sido aislada de una amplia variedad de mamíferos (humanos, animales domésticos y silvestres) y de garrapatas por todo el mundo, incluso de muestras ambientales contaminadas por bacterias excretadas por hospedadores infectados (Maurin y Raoult, 1999; Angelakis y Raoult, 2010; Piñero et al., 2014). Adicionalmente, endosimbiontes parecidos a C. burnetii (‘Coxiella-like endosymbionts’ en inglés) están presentes en garrapatas, sobre todo de la familia Argasidae e Ixodidae (Reeves et al.,

2006; Duron et al., 2014; Machado-Ferreira et al., 2011; Davoust et al., 2014). Bacterias similares a Coxiella han sido descritas como la causa de un cuadro clínico severo observado en aves tropicales mantenidas en cautividad; la infección se ha descrito habitualmente con cuadro mortal (Shivaprarad et al., 2008; Woc-Colburn et a., 2008).

Coxiella burnetii también ha sido detectada en otros artrópodos hematófagos, como las pulgas (Psaroulaki et al., 2014a).

Avances recientes en la investigación sobre Coxiella burnetii y fiebre Q

Durante los últimos años el conocimiento de la fiebre Q en humanos y animales domésticos ha tenido una gran expansión, en parte debido a las constantes noticias de casos humanos en zonas endémicas (Alonso et al., 2015) y por los brotes epidémicos en países como Croacia, Estados Unidos, Guayana Francesa, Holanda, Hungría o Polonia

(Davoust et al., 2014; Morroy et al., 2015; Kersh et al., 2013; Szymańska-Czerwińska et al., 2015; Gyuranecz et al., 2015; Medic et al., 2005). Los avances en salud pública incluyen, entre otros, mejor conocimiento de la epidemiología de la enfermedad (Million y Raoult, 2015), el reconocimiento del papel que juegan los factores del hospedador en la expresión de la fiebre Q aguda y la evolución en el curso de las infecciones crónicas, patología e inmunidad de la infección, desarrollo de protocolos prolongados de administración de antibioterapia frente a la endocarditis producida por C. burnetii

(Million y Raoutl, 2015) y progreso en la prevención a través del desarrollo de vacunas y la mejora de los protocolos de vacunación (Isken et al., 2013; Million y Raoult, 2015).

En sanidad animal los avances han sido, entre otros, el progreso en la comprensión de los factores de riesgo que determinan el mantenimiento y transmisión de la infección (Brom et al., 2015), mejor conocimiento de los factores del huésped y el rebaño que modulan el impacto de la infección (Piñero et al., 2014), un mejor conocimiento sobre la importancia de la infección en la producción ganadera (Oporto et al., 2006, García-Ispierto et al., 2014;

Berri et al., 2005) y mejora de las vacunas y protocolos de vacunación (Roest et al.,

2013a). Sin embargo este impulso significativo en el conocimiento de C. burnetii no se ha visto reflejado en la misma medida en la fauna silvestre, donde los principales avances han consistido principalmente en sacar a la luz el papel de estas especies en el ciclo de vida del patógeno. Sin embargo, estos avances sólo muestran la ‘punta del iceberg’ del potencial de la fauna silvestre en el mantenimiento y la transmisión de C. burnetii. Queda por lo tanto, una labor muy árdua que llevar a cabo en el estudio de este patógeno en las especies silvestre para clarificar su papel en la ecología de C. burnetii, estimar la extensión y la dimensión del riesgo para los propios animales silvestres, los animales domésticos y el ser humano, y para diseñar estrategias de prevención y control del patógeno en la fauna silvestre cuando éste tenga un impacto significativo.

3. Impacto de la fiebre Q en la salud pública, la sanidad animal y la economía

Las consecuencias clínicas de la infección por C. burnetii en humanos y animales llevan consigo importantes costes económicos asociados. Aunque el impacto de la fiebre Q en la salud humana se conoce desde que fue descrito en 1935 (Derrick, 1937), la mayor relevancia de la enfermedad en salud pública se ha puesto en evidencia a partir de los brotes, algunas veces masivos, que han ocurrido a finales del siglo XX y principios del siglo XXI in Bulgaria, Croacia, Francia, Guayana Francesa, Israel, Hungría, Paises Bajos,

Estados Unidos y España (Dupont et al., 1992; Panaiotov et al., 2009; Amitai et al., 2010;

Bjork et al., 2014; Gyuranecz et al., 2014; Davoust et al., 2014; Medic et al., 2005; Alonso et al., 2015). Su impacto económico es grande aunque infravalorado debido a su poco conocimiento y la escasa declaración de casos (Fernández Guerrero, 2014).

Impacto en la salud pública

El incremento mundial del interés por la fiebre Q y su agente causal se pone de manifiesto al observar la tendencia creciente en el número de estudios científicos publicados en las

últimas décadas (Millions et al., 2015) tanto en el ámbito de la salud humana como en el de la sanidad animal. La fiebre Q es rara vez una enfermedad mortal en humanos, pero con frecuencia es una enfermedad debilitante. Actualmente, la fiebre Q es considerada como una enfermedad emergente (Arricau-Bouvery & Rodolakis, 2005). Los seres humanos son altamente susceptibles a la infección por C. burnetii, una sola bacteria es suficiente para desencadenar la infección (Sawyer et al., 1987; Maurin y Raoult, 1999).

Las manifestaciones clínicas en humanos son muy variables, desde casos agudos hasta infecciones crónicas fatales, sin embargo, la mayoría de las infecciones (60%) cursan de forma asintomática, detectándose únicamente la presencia de anticuerpos frente a C. burnetii (Arricau-Bouvery & Rodolakis, 2005; Maurin & Raoult, 1999). La infección aguda, caracterizada por su polimorfismo, presenta manifestaciones clínicas que dependen de la puerta de entrada del patógeno; cursa con mayor frecuencia como un cuadro de neumonía con fiebre elevada (40°C), distrés respiratorio agudo y hallazgos radiográficos inespecíficos. La infección aguda puede manifestarse de manera variable, incluyendo una o varias de las siguientes manifestaciones clínicas o cuadros inflamatorios: fiebre, fatiga, escalofríos, dolor de cabeza, mialgia, erupciones cutáneas, sudoración, náuseas, vómitos, diarrea, tos, dolor de pecho, neumonía, hepatitis, miocarditis, pericarditis, meningoencefalitis e, incluso, la muerte. La proporción de

16 muerte en infecciones agudas tiene una incidencia de entre el 0,9% y el 2,4%

(Kampschreur et al., 2010; Dupont et al., 1992; Parker et al., 2006; Tissot-Dupont &

Raoult, 2008). Un bajo porcentaje de casos agudos, especialmente pacientes con valvulopatías previas, y en menor medida personas inmunodeprimidas y mujeres gestantes, pueden evolucionar a cursos más graves y complicados (crónicos) que pueden presentarse con endocarditis, alteraciones vasculares, procesos osteoarticulares, hepatitis crónica, infecciones pulmonares crónicas y síndrome de fatiga crónica. La incidencia de mortalidad en las infecciones crónicas se situa entre el 1 y el 5% (Maurin & Raoult, 1999).

Sin embargo, en nuestro entorno, se han referido diferentes formas de presentación según el área geográfica: en el norte predominan las neumonías, mientras que en el sur es más frecuente la hepatitis aguda, con hepatomegalia y granulomas (de Alarcón et al., 2003;

Espejo et al., 2014). En la fiebre Q crónica, la manifestación clínica más frecuente es la endocarditis, que se diagnostica, casi exclusivamente, en pacientes con una afección valvular previa, en pacientes trasplantados y en pacientes inmunodeprimidos.

La fiebre Q sigue constituyendo un riesgo laboral principalmente para las personas en contacto con animales domésticos (vacas, ovejas y cabras fundamentalmente) pero también afecta a personas indirectamente relacionadas con animales domésticos (Alonso et al., 2015). La forma más común de contagio es por inhalación de aerosoles que contienen la bacteria (Maurin & Raoult, 1999; Angelakis & Raoult, 2010). De esta forma, dentro de las personas con alto riesgo de sufrir fiebre Q se puede incluir a granjeros, veterinarios, personal de mataderos, personas en contacto con productos lácteos e incluso personas que viven en zonas rurales y personal investigador que trabaje en laboratorios con C. burnetii (Maurin & Raoult, 1999). Los principales reservorios de C. burnetii son los rumiantes domésticos; concretamente, la bacteria es excretada en grandes cantidades mediante secreciones vaginales, leche y heces (Guatteo et al., 2007, Maurin & Raoult,

1999; Angelakis & Raoult, 2010). Aparte de los rumiantes domésticos, otras especies domésticas como perros y gatos pueden ser también fuente de infección para los humanos

(Laughlin et al., 1991; Kosatsky et al., 1984); incluso especies de animales silvestres y artrópodos como las garrapatas son fuente de infección para otros animales y para humanos (González-Barrio et al., 2015a, 2015b – CAPÍTULOS II.1 y II.2; Davoust et al., 2014; Toledo et al., 2009; Kirchgessner et al., 2012a). El vínculo ente fauna silvestre y ser humano en la transmisión de C. burnetii ha sido infravalorado, entre otros motivos, por la dificultad en la trazabilidad de casos humanos con origen en fauna silvestre.

Recientemente se ha especulado sobre este enlace entre fauna silvestre y ser humano

(Schleenvoigt et al., 2015).

Impacto en la sanidad animal

Aparte de la importancia de C. burnetii en salud pública, la fiebre Q en los rumiantes domésticos causa problemas reproductivos, en ocasiones de gran relevancia por el número de pérdidas reproductivas y productivas asociadas. Los problemas reproductivos en rumiantes domésticos incluyen, entre otros, abortos, endometritis e infertilidad. El cuadro clínico y el impacto reproductivo de C. burnetii también puede ser, en cierta medida, extrapolado a la fauna silvestre.

En animales, la fiebre Q en la mayoría de los casos es asintomática. Sin embargo, Coxiella burnetii es uno de los patógenos más importantes causantes de fallo reproductivo en el ganado (Oporto et al., 2006; Agerholm, 2014). Los signos clínicos de la fiebre Q en ruminates domésticos son diversos; C. burnetii ha sido asociada en ganado, principalmente en cabras y ovejas, con casos esporádicos de partos prematuros, abortos y animales nacidos débiles que pueden morir a las pocas horas del parto (Agerholm 2013).

En vacuno, C. burnetii está asociada en su mayoría con problemas de fertilidad

(Agerholm 2014, García-Ispierto et al., 2014). 18

Impacto económico de Coxiella burnetii

Las consecuencias clínicas de la infección en humanos – con brotes masivos como el de los Países Bajos entre 2007 y 2010 (van der Hoek et al., 2012), y las medidas aplicadas para su prevención y control (van Asseldonk et al., 2013) han aumentado el alto, aunque aún infravalorado, impacto económico de C. burnetii. El impacto de la fiebre Q en la salud pública y, por ende en la sociedad y la economía, es significativo (EFSA, 2010).

Las pérdidas en producción en al ganado causan también importantes pérdidas económicas. En la epidemia de los Paises Bajos los costes económicos fueron estimados en 307 millones de euros (Van Asseldonk et al., 2013; 2015), incluyendo: i) costes para el control de la enfermedad en el ganado (testaje, vacunación y sacrificio); ii) costes de las pérdidas productivas en las granjas (pérdida de fertilidad, pérdidas reproductivas, pérdidas en la producción láctea, restricciones reproductivas); iii) costes en salud humana

(hospitalizaciones, tratamientos, pérdida de ingresos económicos por día hospitalizado, probabilidad de síndrome de fatiga crónica, probabilidad de fiebre Q crónica, duración de fiebre Q aguda, años productivos perdidos por enfermedad o fallecimiento, ponderación de las indemnizaciones por discapacidad producida por fiebre Q aguda, ponderación por la discapacidad producida por el sindrome de fatiga crónica, ponderación por la discapacidad producidad por fiebre Q crónica, proporción de bajas en trabajadores con sindrome de fatiga crónica). Otro estudio estima una pérdidas totales causadas en este brote epidémico de fiebre Q en los Paises Bajos de entre 161 y 336 millones de euros

(Tempelman et al., 2011). Morroy et al., (2012) estimó las pérdidas entre 225 y 600 millones de euros. En ambos estudios se estimaron pérdidas en la calidad de vida humana en torno a los 150 millones de euros. Estos ejemplos podrían ser comparable a los costes de brotes similar en otros países desarrollados. Sin embargo, muchos costes asociados a este brote epidémico no fueron incluidos, como los costes en la organización para la

19 gestión del brote y sus causas, así como la financiación científica y cualquier coste potencial sobre la salud de la fauna silvestre y, por ende, la potencial pérdida de biodiversidad y el impacto en los aprovechamientos cinegéticos o de cualquier tipo de la fauna silvestre (turismo, valor intrínseco de la fauna silvestre). La dificultad de estimar los costes económicos ocasionados por la infección por C. burnetii en la fauna silvestre se debe sobre todo a la falta de información sobre los efectos de la infección en la dinámica poblacional en la salud de las poblaciones de fauna silvestre. Estas estimas económicas tampoco han tenido en cuenta los costes producidos por fiebre Q en humanos durante los años previos y siguientes al brote, años en los que los casos de fiebre Q han existido aunque en menor medida. Por lo tanto, costes veterinarios, y sobre todo los costes en la salud pública y las implicaciones sociales parecen haber sido subestimados

(Morroy et al., 2013).

4. Eco-epidemiología de Coxiella burnetii

La epidemiología en la fiebre Q humana está principalmente condicionada por la transmisión vinculada al ganado debido, entre otros factores, a la endemicidad de C. burnetii en el ganado a nivel mundial y al contacto frecuente, directo o indirecto, entre ganado y humanos. La relación que se observa entre la prevalencia de infección por C. burnetii en el ganado y la densidad de animales en las explotaciones (Álvarez et al., 2012;

Piñero et al., 2014) señala que las explotaciones intensivas pueden jugar un papel importante en el riesgo de transmisión a humanos y, por lo tanto, en la aparición de brotes masivos como el de Países Bajos reciente. Con este factor en mente, podríamos hipotetizar que el aumento de brotes de fiebre Q en humanos está probablemente vinculado a los cambios históricos en los sistemas de producción ganadera. Desde el

último cuarto del siglo XX grandes explotaciones intensivas de animales han sustituido a los pequeños sistemas de producción ganadera ligados a zonas rurales, de esta manera,

20 tanto el número de animales como los animales criados por explotación han aumentado considerablemente (Delgado et al., 1999; Thornton, 2010) El incremento en las densidades de animales en las explotaciones intensivas, probablemente debido a que conlleva un aumento en la tasa de interacción entre individuos infectados y susceptibles, favorecería la transmisión y aumentaría la tasa básica de reproducción (R0) del patógeno.

R0 cuantifica el número de individuos susceptibles a los que un individuo infectado es capaza de transmitir la infección. Como ejemplo, el cambio en la producción caprina en los Paises Bajos asociado, entre otros motivos, a la decadencia de la industria porcina en el país debido al control de enfermedades fue el origen del brote masivo de fiebre Q entre

2007 y 2010 (Roest et al., 2011a). Recientemente se observa un aumento en el número de publicaciones de casos de fiebre Q esporádicos en personas que viven en zonas urbanas después de un contacto ocasional con animales de granja o con otros animales domésticos infectados (Laughlin et al., 1991; Kosatsky et al., 1984; Marrie et al., 1988; Marrie, 1996;

Langley et al., 1988), incluso tras contacto con fauna silvestre (Marrie et al., 1986;

Laughlin et al., 1991; González-Barrio et al., 2015c – CAPÍTULO III.2; Davoust et al.,

2014; Schleenvoigt et al., 2015; Eldin et al., 2015). Aunque la tasa de interacción de los seres humanos con la fauna silvestre no es tan elevada como la tasa de interacción con ganado, los patrones actuales de contacto entre la fauna silvestre y el ser humano, tanto directo como indirecto (a través de granjas o fomites), están cambiando (Ruiz-Fons,

2015). Estos cambios se originan debido a la variación en la percepción y el manejo de la fauna silvestre y los alimentos derivados de esta, lo que ha ocasionado la propagación - tanto en la distribución geográfica como en densidad poblacional - de algunas especies de fauna silvestre, como el ciervo rojo (Cervus elaphus), el corzo (Capreolus capreolus), el ciervo de cola blanca (Odocoileus virginianus), el jabalí (Sus scrofa), el zorro rojo

(Vulpes vulpes), la cigüeña blanca (Ciconia ciconia) o el topillo campesino (Microtus

21 arvalis) (Gortázar et al., 2006; Acevedo et al., 2007, 2008; Apollonio et al., 2010, Dawe et al., 2014), entre otros. Cambios en los modos de vida humanos como mayor actividad humana en zonas naturales (turismo rural, deportes al aire libre, avistamientos de fauna), alimentación con productos más naturales (incluyendo productos derivados de la fauna silvestre), crecimiento de las urbanizaciones en zonas naturales, cambios en los aprovechamientos del medio, cambios en los aprovechamientos cinegéticos, e incluso cambios socioeconómicos, pueden conllevar un aumento de la transmisión de patógenos con origen en fauna silvestre (Randolph et al., 2010; Robinsonet al., 2015; Gortázar et al.,

2014a). Además, las cepas de Coxiella burnetii provenientes de la fauna silvestre y que tanto el ganado como los seres humanos no han sido expuestos pueden tener potencial como patógenos emergentes debido a la adquisición de factores de virulencia desconocidos (Gortázar et al., 2014a; Parker et al. 2015). Numerosos patógenos emergentes en humanos tienen su origen en mutaciones genéticas de cepas circulantes en animales silvestres que les han otorgado una gran capacidad de reproducción y transmisión entre humanos tras uno o varios eventos de transmisión de la fauna silvestre al ser humano; podemos citar el virus del Síndrome Agudo Respiratorios (SARS; Sutton

& Subbarao, 2015), el virus del Síndrome Respiratorio del Oriente Medio (MERS; SARS;

Sutton & Subbarao, 2015)o los virus Nipah y Hendra (Daszak et al., 2013; Wood et al.,

2012), entre muchos de los ejemplos existentes.

Por lo tanto, esta sección tiene como objetivo revisar los conocimientos actuales sobre los diferentes aspectos epidemiológicos de Coxiella burnetii en la fauna silvestre a nivel mundial para entender los riesgos potenciales para los animales domésticos y los seres humanos que plantea la fauna silvestre.

Conocimiento actual sobre el estado de Coxiella burnetii en la fauna silvestre

Coxiella burnetii es un patógeno multi-hospedador que es capaz de infectar a un alto número de especies (Maurin and Raoult, 1999). Como muchos patógenos zoonóticos con un amplio espectro de hospedadores, los reservorios silvestres representan un importante riesgo para la salud pública. En este caso el riesgo se presenta a escala global (Kruse et al., 2004), ya que se ha demostrado que C. burnetii puede infectar a un gran número de especies de animales silvestres en diferentes ecosistemas de todos los continentes, desde zonas cálidas como Australia (Potter et al., 2011; Cooper et al., 2013) a zonas tan frías como Alaska (Minor et al., 2013; Myers et al., 2013), incluyendo ecosistemas en islas como Chipre, Japón y Reino Unido (Psaroulaki et al., 2014b; Ejercito et al., 1993;

Meredith et al., 2014), a lugares tropicales y húmedos como la India o la Guayana

Francesa (Yadav et al., 1980; Gardon et al., 2011; Davoust et al., 2014), e incluso en zonas desérticas (Banazis et al., 2010). Muchas especies de mamíferos silvestres, pájaros, peces, reptiles y artrópodos son suscepibles de la infección por C. burnetii (Tabla 1, 2, 3 y 4). La infección también ha sido documentada en fauna silvestre en cautividad, como colecciones zoológicas, safaris o granjas (Tabla 5). En las tablas 1 a 5 se incluye el rango de especies de fauna silvestre y artrópodos en las que se ha analizado la presencia de infección por C. burnetii o evidencias de exposición al patógeno. La mayoría de los estudios sólo han estudiado la fauna silvestre a escalas regionales o locales y muy pocos han intentado proporcionar información a gran escala, por lo que tan sólo tenemos evidencias someras de la situación real de C. burnetii en la fauna silvestre.

Revisando la literaruta científica, 153 especies de 14 órdenes de mamíferos de fauna silvestre han sido analizados para la presencia de anticuerpos y/o presencia de C. burnetii.

Noventa y tres de las 153 especies (60,8%) fueron positivas a la exposición a C. burnetii.

Más de cinco especies han sido analizadas en el orden Artiodactyla (n=31), Carnivora

(n=29), Diprodontia (n=8), Lagomorpha (n=6) y Rodendia (n=61), obteniendo como

23 resultado positivo un 67,7% de las especies de artiodáctilos, un 40,2% de los carnivoros, un 87,5% de los marsupiales diprotodontos, un 100,0% de los lagomorfos y un 60,7% de los roedores. Las técnicas de análisis de la exposición a C. burnetii han sido mayoritariamente técnicas serológicas - ensayo por inmunoadsorción ligado a enzimas

(ELISA), ensayo de inmunofluorescencia (IFA), prueba de la fijación del complemento

(CFT), ensayo de microaglutinación (MAT) y ensayo de aglutinación capilar (CAT) - mientras que la bacteria ha sido aislada o detectada mediantes técnicas moleculares

(Reacción en cadena de la polimerasa, PCR) en 43 de las 91 especies positivas (Tabla 1).

La exposición a C. burnetii ha sido estudiada en 21 órdenes de aves en condiciones de vida libre, incluyendo 154 especies. De estas, 63 especies (40,9%) han sido positivas a C. burnetii. En 10 de los órdenes, más de 5 especies se han estudiado: Accipitriformes

(n=12), Anseriformes (n=8), Charadriiformes (n=10), Columbiformes (n=6),

Falconiformes (n=8), Galliformes (n=8), Gruiformes (n=7), Passeriformes (n=63),

Pelecaneiformes (n=8) y Strigiformes (n=7). En 35 especies de aves de las 63 positivas,

C. burnetii fue aislada o su ADN detectado mediante PCR. El 83,0% de los

Accipitriformes, 50.0% de los Anseriformes, 20.0% de los Charadriiformes, 66.7% de los

Columbiformes, 25.0% de los Falconiformes, 12.5% de los Galliformes, 57,1% de los

Gruiformes, 37,9% de los Paseriformes, 37,5% de los Pelecanieformes y 42,9% de los

Strigiformes fueron positivos a C. burnetii mediantes técnicas serológicas, técnicas moleculares o cultivo celular. En términos generales, el número de especies de aves silvestres analizado es bajo, esto puede indicar que C. burnetii puede estar incluso más extendida en las aves del mundo.

Una de las dos especies de peces en las que se ha estudiado la infección/exposición a C. burnetii fue positiva (Tabla 3).

Otras 19 especies de mamíferos y 7 de aves silvestres en cautividad (zoos, safari, granjas, colecciones personales) fueron positivas a C. burnetii (Tabla 5). Cuatro especies de anfibios (Orden Anura) junto a 5 especies de reptiles (Orden Squamata y Testudines) han sido analizados para la exposición a C. burnetii. Cuatro especies, incluyendo serpientes, lagartos y tortugas, fueron positivas mediante PCR.

Finalmente, en 17 de 27 especies de garrapatas duras - familia Ixodidae – C. burnetii ha sido detectada mediante PCR o cultivo celular (Tabla 5). En tres especies de garraptas blandas - familia Argasidae - y en tres especies de pulgas se detectó C. burnetii por medio de técnicas moleculares (Tabla 4).

A día de hoy, hay muy pocos estudios sobre el estado de C. burnetii y su ciclo en especies de fauna silvestre y/o de vectores artrópodos. La mayor parte de la información ha sido obtenida por estudios parciales u oportunistas con enfoques inapropiados para obtener información representativa. Se debería tender en el futuro a mejorar el conocimiento del estado de C. burnetii en las especies de fauna silvestre, sobre todo aquellas que son abundantes y con una amplia distribución, con aumento en sus tendencias poblacionales y con potencial de interacción medio-alto con humanos y ganado. Estos enfoques permitirán mejorar la comprensión de la ecología de C. burnetii, lo que es esencial para su control y eventual erradicación de las explotaciones ganaderas y las poblaciones humanas.

Factores que modulan la dinámica de Coxiella burnetii en fauna silvestre

Si la información actual a gran escala y a largo plazo sobre el estado de C. burnetii en especies de vida silvestre es escasa, los estudios epidemiológicos que analizan los factores de riesgo para el mantenimiento de este patógeno en la fauna silvestre son aún menos abundantes. En esta sección, y debido a la escasez de estudios, se abordan los factores

25 potencialmente moduladores de la dinámica de transmisión de C. burnetii en la fauna silvestre utilizando para ello evidencias científicas existentes en la fauna silvestre, basándose en el conocimiento previo de los factores que impulsan otros patógenos multi- hospedador y en el conocimiento existente en el ganado y en la especie humana.

Identificar los factores potenciales que modulan el mantenimiento y la transmisión de C. burnetii en la fauna silvestre es esencial para estimar los riesgos y para diseñar y aplicar cualquier potencial método de control (Boadella et al., 2012a). Se analiza el potencial efecto de factores propios de los hospedadores (demografía, ciclo de vida y características fisiológicas del hospedador), factores ambientales (condiciones meteorológicas y climáticas) y factores propios del patógeno (virulencia, diversidad genética).

Factores demográficos. La susceptibilidad de un hospedador silvestre por un patógeno y su capacidad para multiplicar y excretar el patógeno son pre-requisitos para asumir que ese hospedador pueda actuar como reservorio (Wobesser et al., 1994). Sin embargo, para estimar el posible papel como reservorio de un hospedador silvestre para un patógeno concreto debemos conocer la capacidad de dicho hospedador para mantener el patógeno, es decir, que R0 sea ≥1 (Metcalf et al., 2015). Diversos factores relacionados con la dinámica poblacional del hospedador (distribución geográfica, tendencias demográficas, densidad y agregación) pueden condicionar la tasa de reproducción básica del patógeno y, con ello, determinar que este circule o que se extinga de la población.

El riesgo que dos especies de hospedadores competentes para un patógeno compartido con otros animales y con el ser humano representa podría variar en función de su distribución geográfica. Por ejemplo, el rebeco alpino (Rupicapra rupicapra) y el ciervo rojo son ambos susceptibles a la infección por C. burnetii (Pioz et al., 2008a; González-

Barrio et al., 2015a – CAPÍTULO II.1), sin embargo la extensión de sus áreas de distribución son muy diferentes. El primero es un ungulado de alta montaña con unos

26 requerimientos ambientales particulares que lo mantienen restringido a los ecosistemas montañosos del suroeste de Europa (Herrero et al., 2008). Sin embargo, el ciervo rojo presenta una amplia distribución en Europa, norte de África, sur de América y Asia

(Flueck et al., 2003; Ludt et al., 2004). Así, sólo la mera variación en la distribución geográfica de dos especies de hospedadores competentes conllevaría una diferencia en el riesgo que ambas representan para la transmisión de C. burnetii a ganado o ser humano.

Esta hipótesis ha sido postulada para otros patógenos compartidos, como por ejemplo en el virus de la lengua azul (BTV) (Ruiz-Fons et al., 2008a; Ruiz-Fons et al., 2014a). La prevalencia de BTV fue ligeramente más alta en gamo (Dama dama) que en ciervo rojo en el sur de España, lo que a priori señalaría un mayor riesgo de transmisión por parte del gamo en comparación con el ciervo. Sin embargo, a una escala mayor, la distribución geográfica más restringida del gamo señalaría que su papel en la transmisión de BTV es más limitado que el del ciervo. Aunque se debe señalar que el riesgo no debería medirse sólo en base a rangos de distribución geográfica actuales de los hospedadores silvestres, sino en las tendencias de distribución geográfica que estos presentan y que determinarán el riesgo en un futuro próximo. Actualmente tenemos evidencias de que algunas especies competentes para C. burnetii presentan unas tendencias geográficas crecientes a gran escala, como el ciervo rojo (Apollonio et al., 2010), el corzo (Acevedo et al., 2005;

Acevedo et al., 2010a), el ciervo de cola blanca (Gallina et al., 2008) o el jabalí (Massei et al., 2015; Schöning et al., 2013). Estas especies representarían un mayor riesgo como fuente de C. burnetii para el ganado y el ser humano que otras especies silvestres con tendencias de distribución geográfica estables o decrecientes como el rebeco alpino

(Rupicapra rupicapra) y el rebeco pirenaico (Rupicapra pyrenaica) (Herrero et al., 2008;

Aulagnier et al., 2008a), o la cabra montés (Capra hispanica) y el íbice alpino (Capra ibex) (Herrero y Pérez, 2008; Aulagnier et al., 2008b). Desafortunadamente, la

27 información existente sobre las tendencias geográficas actuales de los mamíferos y reptiles silvestres son escasos. Por el contrario, la información de tendencias de especies de aves silvestres en los países desarrollados es monitorizada por algunas organizaciones no gubernamentales (SEO/Bird life; WWF).

R0 es, por definición, una variable que depende de la tasa de interacción entre individuos infectados y susceptibles (Dobson & Foufopoulos, 2001). A mayor tasa de interacción más alto es el riesgo de transmisión. Las interacciones entre los individuos de una especie dependen de varios factores, incluyendo su dinámica poblacional, factores conductuales de los hospedarores y factores ambientales (Dobson & Foufopoulos, 2001). Centrándonos en la dinámica de población del hospedador, las interacciones entre individuos son más frecuentes a altas densidades poblacionales que cuando la densidad es baja (Gortázar et al., 2006). Este factor por sí mismo puede condicionar R0 y determinar el mantenimiento o la extinción del patógeno en la población (ej., Rossi et al., 2005a, 2005b en la transmisión del virus de la peste porcina clásica en jabalí). Estudios en vacuno de leche encontraron que la densidad del hospedador tenía efecto en el riesgo de exposición a C. burnetii (Álvarez et al., 2012; Piñero et al., 2014). Un estudio reciente en conejo de monte

(Oryctolagus cuniculus) en la península ibérica encontró que los valores de seroprevalencia frente a C. burnetii más altos se daban en poblaciones de conejo gestionadas con fines cinegéticos, hecho que promueve altas densidades (González-

Barrio et al., 2015b - CAPÍTULO II.2). Los efectos de la densidad no se aplican

únicamente a una poblacion de hospedadores en particular, sino que también se puede aplicar para comparar el riesgo potencial de diferentes hospedadores competentes. Por ejemplo, tanto el ciervo rojo como el corzo están ampliamente distribuidos en Europa

(Apollonio et al., 2010). Sin embargo, mientras el ciervo rojo alcanza densidades de hasta

70 individuos/Km2, en el corzo las densidades más altas registradas en Europa no

28 sobrepasan los 7 individuos/Km2 (Acevedo et al., 2008; Prokešová et al., 2006; Walander et al., 2012). Por lo tanto, es esperable que la tasa de interacción entre ciervos y, por lo tanto R0, sean mayores en el ciervo que en las poblaciones de corzo. Sin embargo, como

C. burnetii es un patógeno transmitido principalmente de forma indirecta y con una amplia gama de hospedadores (Maurin & Raoult, 1999), otros factores, discutidos en los siguientes apartados, deberían considerarse para estimar la relevancia de la densidad del hospedador en las tasas de interación y en la transmisión de C. burnetii. De hecho, la prevalencia en las poblaciones de corzo en los Paises Bajos, con densidades de 4 individuos/Km2 (Montizaan & Siebenga, 2010), fue más alta (23,0%) que la media de la seroprevalencia en ciervos silvestres en la península ibérica (3,6%; González-Barrio et al., 2015a - CAPÍTULO II.1) en las que las densidades de ciervo pueden alcanzar valores de hasta 70 individuos/Km2 (Acevedo et al., 2008). Otros factores aparte de la densidad, teniendo en cuenta los sesgos potenciales presentes en los estudios existentes (González-

Barrio et al., 2015a - CAPÍTULO II.1), pueden modular la tasa de transmisión de C. burnetii. La densidad poblacional está en parte condicionada por la capacidad reproductiva - prolificidad - del hospedador, y este parámetro podría indirectamente modular la dinámica de C. burnetii. Especies con alta prolificidad, como el conejo de monte (Dekker et al., 1975) o el jabalí (Ruiz-Fons et al., 2006), pueden incrementar su densidad poblacional en poco tiempo y proporcionar un gran número de animales susceptibles a la infección por C. burnetii, manteniendo de esta manera el patógeno en la población. Actualmente, muchas de las herramientas disponibles para la estima de densidad poblacional de especies de fauna silvestre no son aplicables a las diferentes regiones en las que estas especies están presentes (Lancia et al., 1994; Acevedo et al.,

2007), y el esfuerzo necesario para realizar estudios sobre la relación entre la densidad y el riesgo de la exposición a C. burnetii es muy grande, lo que condiciona la viabilidad de

29 estimar a gran escala el potencial de la densidad del hospedador en la dinámica de C. burnetii. Otro parámetro que condiciona la tasa de interacción entre individuos en una población es la agregación de los individuos (Acevedo et al., 2007). Esta depende principalmente de la gestión de la fauna silvestre, de las características comportamentales de la especie y de factores ambientales, por lo que se discute más adelante en las secciones correspondientes.

La particular amplia gama de hospedadores de C. burnetii quizás dificulta la comprensión de los factores que determinan su transmisión debido a la influencia de la composición de la comunidad de hospedadores en dicha transmisión. El efecto de la coexistencia de hospedadores competentes y no competentes en el mantenimiento de C. burnetii es poco conocido, incluso en rebaños mixtos de rumiantes domésticos. Haydon et al. (2008) proponen una serie de escenarios con variaciones en la comunidad de hospedadores que pueden modular el mantenimiento y transmisión de patógenos multi-hospedador. Estos escenarios no sólo contemplan el papel de hospedadores competentes si no también el efecto de especies no competentes en simpatría. Para entender este efecto en el caso de

C. burnetii podemos utilizar el ejemplo del papel de la presencia de garrapatas en una comunidad de hospedadores competentes y no competentes. Aunque actualmente el papel de las garrapatas en el mantenimiento y transmisión de C. burnetii no está claro, cualquier papel de las garrapatas se vería reforzado por hospedadores que, a pesar de no ser susceptibles a la infección por C. burnetii, tuviesen un papel importante en el mantenimiento de las poblaciones de garrapatas. El efecto de hospedadores no competentes para el patógeno pero con un papel en la dinámica de vectores competentes se ha demostrado en la tranmisión de Borrelia burgdorferi, agente causal de la enfermedad de Lyme (Biesiada et al., 2012). Los ungulados silvestres son hospedadores clave en el mantenimiento de los vectores de B. burgdorferi, principalmente Ixodes

30 ricinus en Europa e I. scapularis en Norteamérica, y , por lo tanto, en el mantenimiento y transmisión de este patógeno a pesar de que son incompetentes para B. burgdorferi por sí mismos (Cook et al., 2014). La coexistencia de hospedadores competentes para B. burgdorferi (principalmente pequeños mamíferos y aves terrestres) con hospedadores clave para sus vectores aumenta el riesgo de transmisión a terceras especies (Baneth,

2014). El efecto de la variación en la comunidad de hospedadores en la dinámica y transmisión de C. burnetii es actualmente desconocido. Sin embargo, estudios recientes de epidemiología molecular sugieren que los genotipos de C. burnetii que circulan en cada especie de hospedador difieren de aquellos que circulan en hospedadores en simpatría (CAPÍTULO II.4). Los corzos analizados mediante MLVA en Holanda estaban infectados por genotipos de C. burnetii diferentes de los genotipos existentes en el ganado con el que coexistían durante el brote de 2007-2010 (Rijks et al., 2011).

Recientemente, el análisis genético mediante MLVA de genotipos presentes en conejo y ciervo en simpatría revela que los genotipos son más similares dentro de cada hospedador que entre hospedadores. A pesar de que ciervo y conejo en este estudio comparten pastos, los genotipos compartidos por ambas especies son escasos (CAPÍTULO II.4).

Resultados similares se han obtenido al analizar los genotipos que circulan en ovejas y vacas dentro de un mismo rebaño (de Bruin et al., 2012). Estos hallazgos sugieren que podrían existir relaciones hospedador-patógeno específicas causadas por adaptaciones hasta ahora desconocidas de C. burnetii a determinadas características del hospedador.

Sin embargo, la información es todavía escasa y esta hipótesis debe ser probada aumentando del número de genotipos y mediante el empleo de diferentes técnicas de tipado molecular, quizás a través de secuenciación masiva de todo el genoma de C. burnetii (Pearson et al 2014; Massung et al., 2012).

Factores individuales del hospedador. Tanto el patógeno como el hospedador interactuan entre sí y esta interacción puede ser modulada tanto por factores del patógeno (que serán objeto de estudio más adelante) como por factores individuales del hospedador

(comportamiento y características fisiológicas). El gregarismo es un comportamiento particular que puede promover la exposición a patógenos de individuos susceptibles en poblaciones donde los patógenos están presentes (Lee et al., 2008). De este modo, especies gregarias como los bóvidos silvestres - arruí (Ammotragus lervia), muflón (Ovis aries musimon), íbice alpino, rebeco alpino/pirenaico, diversas especies de gacelas, algunos cérvidos - ciervo rojo y gamo, algunos carnívoros - como el león (Panthera leo) o el lobo (Canis lupus), o algunas aves que forman colonias - cigüeña blanca (Ciconia ciconia), estorninos o buitres, podrían hipotéticamente estar sujetos a exposiciones más elevadas a C. burnetii cuando las comparamos con especies solitarias - algunos antílopes, corzo, alce (Alces alces), zorro rojo (Vulpes vulpes) o la mayoría de aves rapaces, entre otros. Podríamos considerar que el gregarismo puede no ser homogéneo para cada individuo dentro de una especie hospedadora concreta. Como ejemplo, las hembras de jabalí son gregarias y viven en grupos familiares de tamaño variable con sus crías, en contraste a los machos adultos que son solitarios y solo ocasionalmente forman grupos de pequeño tamaño con otros machos (Fernández-Llario and Carranza, 2000). Por lo tanto, los patrones de comportamiento sesgados por sexo y/o edad pueden resultar en tasas de exposición variables de los individuos dentro de una especie hospedadora (Ruiz-

Fons et al., 2013). El efecto del gregarismo en la dinámica de C. burnetii es poco conocida y los datos existentes son escasos, incompletos y controvertidos para apoyar la hipótesis propuesta. El ejemplo del corzo en los Paises Bajos y del ciervo rojo en España podría contradecir esta hipótesis ya que el corzo forma grupos muy reducidos (de 2 a 5 individuos, Pays et al., 2007) en contraste con el ciervo rojo (grupos de más de 60-80

32 individuos, Clutton-Brock et al., 1982). Así mismo, especies en las que su comportamiento es solitario como el perezoso (Bradypus trydactilus) en Cayena,

Guayana Francesa (Davoust et al., 2014) ha sido documentado como un potencial reservorio de C. burnetii. Por lo tanto, estudios epidemiológicos específicos deben ser diseñados en el futuro para poner a prueba esta hipótesis en diferentes hospedadores silvestres.

Alimentarse puede incluso condicionar hipotéticamente la exposición a C. burnetii. Las especies susceptibles en el último eslabón de la cadena alimentaria (depredadores grandes y medianos) deberían presentar un mayor riesgo de exposición a C. burnetii a través de la exposición a animales infectados y/o sus cadáveres. Dos aves carroñeras, el buitre leonado (Gyps fulvus) y el milano negro (Milvus migrans), han mostrado prevalencias de infección por C. burnetii más altas que especies de ungulados, carnívoros y lagomorfos con los que coexisten en el norte de España (Astobiza et al., 2011a). Aunque el zorro rojo en este mismo estudio no mostró ninguna evidencia de infección por C. burnetii, otros estudios en Estados Unidos y Reino Unido encuentran altas prevalencias (40-60%; Tabla

1) de anticuerpos frente a C. burnetii en este carnívoro (Enright et al., 1971; Willeberg et al., 1980; Meredith et al., 2014). Recientemente, Cumbassá et al., (2015) describen genotipos mediante MLVA en meloncillo (Herpestes ichneumon) en Portugal.

Curiosamente, estos genotipos difieren de otros genotipos descritos en el ganado y en humanos en Portugal, pero son más similares a los genotipos tipados con la misma técnica que han sido descritos en conejo de monte en España (CAPÍTULO II.4). El conejo de monte constituye la principal presa del meloncillo en la peninsula Ibérica (Delibes et al.,

1984). Estos datos sugieren que C. burnetii podría utilizar las relaciones depredador-presa en su favor.

Los factores comportamentales de los individuos podrían modular la exposición a C. burnetii, pero otros factores del individuo podrían también modular la infección una vez se ha producido la exposición al patógeno. El conocimiento sobre el efecto de la capacidad inmunológica del hospedador en modular la infección por C. burnetii es actualmente escaso y la mayor parte de la información proviene de experimentos en laboratorio con pequeños animales como modelo (Bewley, 2013). La capacidad inmunológica del hospedador modula la infección por cualquier patógeno, por lo que cualquier factor que condicione la capacidad inmunológica del hospedador podría modular la infección por C. burnetii en el organismo. En fauna silvestre la capacidad inmunológica de un hospedador depende en gran parte de su condición física, que en

última instancia depende de la disponibilidad y calidad de alimento, de la disponibilidad de agua, y del estrés (Moller et al., 1998; Coop & Kyriazakis, 1999, 2001; Lochmiller &

Deeremberg, 2000; Fernández-de-Mera et al., 2009). Por lo tanto, el estado físico general de los hospedadores dentro de una población podría, en teoría, modular la dinámica de la infección por C. burnetii. La inmunidad adquirida pasiva, por ejemplo, derivada de la transmisión de anticuerpos de la madre a la cría (anticuerpos maternales), podría tener un papel en la modulación de la infección por C. burnetii. Los mamíferos recien nacidos adquieren pasivamente anticuerpos maternales frente a C. burnetii de sus madres durante las primeras etapas de la lactancia (Tutusaus et al., 2013). Aunque el papel de los anticuerpos derivados de la madre en la protección contra la infección no ha sido probado en los rumiantes domésticos, recientes evidencias en ciervo rojo sugieren que las crías están protegidas contra la infección de C. burnetii en sus primeros meses de vida como consecuencia de la elevada dosis de anticuerpos proporcionados en la leche materna

(González-Barrio et al., 2015d - CAPÍTULO II.3). Por otro lado, la inmunidad global de la población y su cambio en el tiempo podrían modular la dinámica de C. burnetii en

34 situaciones endémicas, determinando variación temporal en el estado de C. burnetii

(prevalencia y presión de infección) en las poblaciones. Esto ha sido recientemente propuesto para explicar la dinámica temporal variable observada en rumiantes tanto domésticos (Piñero et al., 2014) como silvestres (González-Barrio et al., 2015c –

CAPÍTULO II.3).

Factores ambientales. A pesar de la alta capacidad de resitencia ambiental de C. burnetii, condiciones meteorológicas como la humedad, la temperatura, la velocidad del viento, e incluso la composición del suelo (contenido de materia orgánica, estructura del suelo, contenido de humedad) podrían modular la supervivencia de C. burnetii en el medio y con ello la transmisión entre hospedadores. Sin embargo, el potencial efecto de las condiciones ambientales sobre la superviviencia y transmisión de C. burnetii es aun escaso (Pascual-Velasco, 1996) aunque se conoce el efecto de ciertos condicionantes ambientales sobre la supervivencia de C. burnetii (Maurin & Raoult, 1999). La transmisión mediante aerosoles es la forma de transmisión de C. burnetii documentada más efectiva. La formación y la dispersión de aerosoles conteniendo C. burnetii podría ser teóricamente modulada por la humedad del aire (indirectamente relacionada con la temperatura del aire), por el tipo de suelo (tamaño de la partícula del suelo) o por la velocidad del viento, entre otros factores. El viento puede provocar la dispersión de C. burnetii a largas distancias (Tissot-Dupont et al., 2004; Dorco et al., 2012; O'Connor ET

AL., 2015), pero la formación de partículas infectadas cargadas con formas infectantes de C. burnetii podría verse afectada por factores como la humedad o el propio suelo. No existen, o son escasas, las pruebas científicas que analicen el efecto de las condiciones ambientales sobre la transmisión de C. burnetii. Variaciones en la temperatura media de primavera resultaron constituir un factor de riesgo para el riesgo de exposición del ciervo rojo a C. burnetii en la península ibérica (González-Barrio et al., 2015a – CAPÍTULO

II.1). Los autores hipotetizan que un efecto indirecto en el aumento de temperatura media en primavera podría modular la transmisión debido a la existencia de mejores condiciones para la formación de aerosoles que contengan la bacteria durante la principal época de excreción descrita en ciervo rojo (González-Barrio et al., 2015d – CAPÍTULO II.3).

Factores del patógeno. Las relaciones hospedador-patógeno son dependientes tanto de factores del hospedador como del patógeno. Los patógenos han evolucionado con sus hospedadores a lo largo de milenos, y estos les han permitido adquirir o desechar aquellas características que favorecen o disminuyen, respectivamente, sus habilidades para replicarse en los hospedadores. La diversidad de cepas de C. burnetii que circulan en todo el mundo debe ser tan alta como su gama de hospedadores. Sin embargo, la información sobre la diversidad genética de C. burnetii en todo el mundo es reciente e incompleta

(Piñero et al., 2015; Santos et al., 2012; Tilburg et al., 2012a) y, por lo tanto, es necesaria mucha más información de la existente para caracterizar la diversidad de cepas y variantes patogénicas existentes e identificar potenciales factores de virulencia en ellas.

Factores que favorecen la transmisión de Coxiella burnetii en la interfaz fauna silvestre- ganado doméstico-humano

El grado de interacción de la fauna silvestre con el ganado y/o el ser humano (tanto directo como indirecto) es un parámetro clave para estimar el riesgo de exposición de C. burnetii desde la fauna silvestre a hospedadores que son de nuestro interés. Dado que las tasas de interacción serían extremadamente variables dependiendo de la composición de la comunidad de hospedadores, de la demografía, de las condiciones ambientales, etc., en esta sección se analiza el efecto potencial de aquellos factores que pueden dar lugar a un incremento de la interacción entre la fauna silvestre, el ganado y el ser humano que potencialmente podrían llevar a la transmisión inter-específica de C. burnetii.

Los factores de riesgo que pueden dar lugar al aumento en la transmisión de C. burnetii dentro de las poblaciones de especies de fauna silvestre - que han sido analizados en la sección anterior - podrían consecuentemente aumentar el riesgo de transmisión a ganado y humanos. Sin embargo, otros factores propios del ganado y de los humanos (sistemas de producción, patrones de comportamiento, etc.) pueden también actuar como factores de riesgo para la transmisión de C. burnetii desde la fauna silvestre. Actualmente existe poca información sobre la fuente de origen de muchos casos de fiebre Q en humanos

(EFSA 2010). Estudios epidemiológicos apropiados sólo se llevan a cabo después de los brotes de fiebre Q que implican a varias personas geográficamente relacionadas (de Bruin et al., 2012; Tilburg et al., 2012a, Tilburg et al., 2012b; Sulyok et al., 2014) y estos están vinculados principalmente a la ganadería debido a la proximidad de las instalaciones ganaderas y las poblaciones humanas. Sin embargo los casos relacionados con la fauna silvestre son escasos, esporádicos y de relevancia local, y estos no son seguidos desde el origen con investigaciones moleculares en seres humanos y fauna silvestre (Laughlin et al., 1991; Eldin et al., 2015; Schleenvoigt et al., 2015). Este hecho probablemente viene en gran parte condicionado por la percepción que los responsables de la salud pública tienen sobre el papel de la fauna silvestre en la transmisión de C. burnetii a humanos. Sin embargo, investigadores con una amplia visión de conjunto de los posibles riesgos asociados a la fauna silvestre han sugerido - aunque no demostrado – que el origen de varios casos de fiebre Q humana estaba en la fauna silvestre (Marrie et al., 1986; Laughlin et al., 1991; Davoust et al., 2014; Schleenvoigt et al., 2015). Hasta la fecha, desafortunadamente, ningún caso de fiebre Q en humano con un origen probado de la fauna silvestre ha sido documentado a pesar de que los mismos genotipos han sido encontrados en humanos y animales silvestres (CAPÍTULOS II.4 y II.5). De todas formas, cualquier factor que aumente la interacción entre la fauna silvestre y los seres

37 humanos constituiría potencialmente un factor de riesgo para la transmisión de C. burnetii. A continuación se analiza una serie de factores que pueden conducir a un aumento en las interacciones directas e indirectas entre fauna silvestre y ser humano y que podrían constituir un riesgo para la transmisión del patógeno.

Cambios en los sistemas de producción del ganado doméstico. Los sistemas de producción del ganado han cambiado considerablemente en las últimas décadas practicamente en todo el mundo, pero especialmente en paises desarrollados (Thornton,

2010). Los cambios han consistido en el aumento del número de animales reproductores por granja (Hansen et al., 2014) pero la demanda de productos más ‘naturales’ y

‘ecológicamente sostenibles’ podría conllevar un aumento de la ganadería en condiciones extensivas. La intensificación de la ganadería ha sido el único camino a seguir para proveer de suficiente alimento a la población humana en exponencial crecimiento

(Thornton, 2010; Davis & D'Odorico, 2015; Delgado et al., 2005). Por otra parte, la ganadería extensiva se ha visto incrementada notablemente como consecuencia de la percepción de los consumidores como mucho más sostenible, natural y como manera más biológica de la obtención de alimentos de origen animal (Karesh et al., 2005) y debido a la creciente preocupación y la legislación sobre bienestar animal (Fraser et al., 2014).

Estos cambios pueden tener implicaciones en la dinámica de los patógenos por el incremento en la transmisión dentro del rebaño de patógenos (en explotaciones intensivas con altas densidades) o por el incremento en las enfermedades compartidas entre fauna silvestre y ganado doméstico (en explotaciones ganaderas en extensivo). La ganaderia en extensivo puede reducir tanto el impacto del medio ambiente como los costes económicos asociados en términos de suplementación de alimento ya que los animales pueden aprovechar los recursos naturales; por ejemplo la alimentación a base de bellotas y pasto del cerdo ibérico en dehesas en la península ibérica (Rodriguez-Estevez et al., 2009). Sin

38 embargo, la ganadería en extensivo podría conllevar problemas en términos de salud de los animales, ya que conlleva una mayor tasa de interacción con la fauna silvestre (Olea

& San Miguel-Ayanz, 2006; Gortázar et al., 2010) en la que el control de patógenos es nulo o escaso. Existen numerosos ejemplos de enfermedades compartidas entre fauna silvestre y ganado doméstico: i) Brucelosis transmitida desde íbices a vacas en los alpes franceses (Mick et al., 2014); ii) Virus de la enfermedad de Aujeszky transmitido desde jabalí a cerdo doméstico producido en extensivo en Francia (Hars et al., 2009); o iii)

Transmisión de peste porcina africana entre jabalí y cerdos de traspatio en Sicilia y en el este de Europa (Laddomada et al., 2000; Sánchez-Vizcaíno et al., 2015). En el caso de C. burnetii el riesgo de transmisión entre fauna silvestre y ganado también se incrementa con el aumento del grado de interacción asociado a la producción de ganado en extensivo.

Sin embargo, la transmisión aerógena indirecta de este patógeno conlleva que el riesgo de transmisión entre fauna silvestre y ganado no esté restringido únicamente a la ganadería en extensivo. Cualquier vínculo entre la fauna silvestre y el ganado que favorezca la transmisión de C. burnetii constituiría además un vínculo indirecto que favorecería la transmisión indirecta a humanos desde la fauna silvestre mediada por el ganado.

Cambios en los usos del suelo. El rápido e imparable desarrollo de la población humana

(Worldometers. [http://worldometers.info/]) promueve la ocupación de los espacios naturales y la explotación de sus recursos con la consiguiente perturbación de los ecosistemas. El incremento de la población humana incrementa la demanda de proteínas

(Karesh et al., 2005) promoviendo la sobreabundancia de algunas especies. Cambios recientes en los usos agrícolas del suelo en Castilla y León - transformación de una agricultura de secano a una agricultura de regadío - están detrás del aumento de las poblaciones de topillo campesino (Microtus arvalis) y del aumento de la prevalencia de

Francisella tularensis (Jareño et al., 2015). Los cambios en los planes agrícolas de la

Unión Europea en las últimas décadas con incrementos en la producción de los cultivos de regadío, junto con políticas de protección medioambientales, el aumento de la cubierta vegetal (Gortázar et al., 2000) y el aumento de las temperaturas medias invernales, han representado el aumento masivo en la distribución del jabalí y su densidad en toda Europa

(Apollonio et al., 2010), y con ello han incrementado la preocupación actual sobre la propagación del virus de la peste porcina africana y otras enfermedades compartidas entre jabalí y ganado (Sánchez-Vizcaíno et al., 2015). En estos escenarios de aumento de especies silvestres particulares, la interacción con humanos, animales domésticos y otras especies de fauna silvestre es más frecuente. El efecto de estos cambios en la situación actual de C. burnetii en la fauna silvestre es desconocida y por lo tanto la predicción de cualquier efecto futuro es difícil y completamente especulativa.

Cambios en los modelos de gestión cinegética. La industria cinegética está aflorando en los países desarrollados y subdesarrollados de todo el mundo (Apollonio et al., 2010;

Lindsey et al., 2012). La caza promueve la preservación de las especies de interés cinegético en detrimento de especies simpátricas y, por lo tanto, promueve el aumento de la densidad de las primeras. Esta es una de las principales causas de aumento de ungulados cinegéticos en Europa (Hagen et al., 2014; Hartley & Gill, 2010; Diaz-Fernandez et al.,

2013; Apollonio et al., 2010). La gestión que llevan a cabo los promotores cinegéticos pueden jugar un importante papel en la transmisión de patógenos en las poblaciones de fauna silvestre y, por ende, a ganado y humanos (Gortázar et al. 2006). La introducción de alimentación suplementaria para aumentar las bolsas de caza ha perturbado de manera notable la regulación natural ejercida por la capacidad de carga del medio sobre la fauna silvestre. Así mismo, la suplementación de alimento incrementa la agregacion de los individuos en los puntos de alimentación, lo que promueve un aumento de R0 y de las

40 prevalencias de infección (Gortázar et al., 2010; Alexander et al., 2010). A pesar de esto, la gestión de la caza no se ha encontrado como un factor de riesgo para la exposición del ciervo rojo a C. burnetii (González-Barrio et al., 2015a – CAPÍTULO II.1), ya que la seroprevalencia fue mayor en poblaciones no gestionadas que en aquellas con algún sistema de gestión cinegética (vallado, alimentación suplementaria y/o suministro de agua). Las actividades cinegéticas intensivas también pueden promover la translocación de animales entre poblaciones para introducir mejoras en la calidad de los trofeos, par evitar la consanguinidad o para reforzar las poblaciones (Apollonio et al., 2010; Lyons et al., 2013). Desafortunadamente, a pesar de las regulaciones sobre el transporte de animales cinegéticos en los países desarrollados, muchos desplazamientos se hacen de manera ilegal. Esto representa un aumento del riesgo de introducción de patógenos exóticos en zonas potencialmente susceptibles con alta densidad de hospedadores, elevadas agregaciones y la altas tasas de interacción con el ganado y los seres humanos.

En algunos terrenos privados coexisten los aprovechamientos cinegéticos y ganaderos, lo que promueve la interacción entre la fauna silvestre y el ganado, y así el intercambio de patógenos compartidos (Delibes-Mateos et al., 2009; Kukielka et al., 2013). Actualmente, el comercio de fauna silvestre es uno de los principales problemas para la transmisión de agentes infecciosos entre especies (Swift et al., 2007; Gómez et al., 2008; FAO). El efecto de la gestión con fines cinegéticos en el dinámica de C. burnetii en fauna silvestre debería ser objeto de estudio en futuros trabajos de investigación.

5. Patogénesis y transmisión

La forma en la que la infección por Coxiella burnetii se produce en el organismo está bien definida tanto en animales dométicos (Angelakis y Raoult, 2010) como en humanos

(Maurin y Raoult, 1999), pero casi nada se sabe de las particularidades de los diferentes hospedadores silvestres en la forma en que la infección por C. burnetii se establece, cómo

41 se replica, cómo se excreta y cómo se transmite. Por lo tanto, aunque se asimile que lo que se conoce de la patogenia en animales domésticos es válido para la fauna silvestre, seguramente existen mecanismos particulares del hospedador que modulan la infección por C. burnetii. La información actual existente sobre estos factores es muy escasa.

La principal forma de contraer la infección por C. burnetii en animales domésticos y humanos es a través de aerosoles (Badubieri, 1959). Otras vías de infección - por ejemplo, oral y reproductiva - son consideradas una alternativa poco común para adquirir la infección, aunque son posibles (Maurin & Raoult, 1999; Ruiz-Fons et al. 2014b;

González-Barrio et al., 2015e – CAPÍTULO III.1). Por lo tanto, las vías respiratorias serían teóricamente el primer lugar de interacción hospedador-patógeno por transmisión aerógena en fauna silvestre. En humanos, el período de incubación de fiebre Q aguda puede variar de 2 a 6 semanas después de la exposición, aunque la infección permanece asintomática en mas de 50% de los casos, dependiendo de la dosis de C. burnetii (Maurin

& Raoult, 1999). En animales este período de incubación es variable. La dosis infectiva para C. burnetii puede ser tan baja como un solo organismo inhalado (Van schaik et al.,

2013). En humanos, la enfermedad sintomática suele durar 1-2 semanas (Van schaik et al., 2013). En los casos sintomáticos, la enfermedad aguda se resuelve espontáneamente en 1-6 semanas y se mitiga eficazmente por tratamiento con antibióticos como la tetraciclina y las cefalosporinas de tercera generación (Million et al., 2010; Raoult et al.,

1999). Algunos estudios han estimado que en el 1-5% de los casos, la fiebre Q aguda desarrolla a una infección crónica (Marmion et al., 1996; 2005). Neumonía y problemas cardíacos se han descrito también en vacuno (Maurin & Raoult, 1999; Guatteo et al.,

2006; Saegerman et al., 2011).

Una vez inhalada, o ingerida, la forma extracelular de Coxiella burnetii (SCV) se adhiere a la membrana de los macrófagos locales y se internaliza en las células del hospedador.

La primera replicación de C. burnetii se produce en los ganglios linfáticos regionales de la principal vía de transmisión, por lo general en la zona orofaríngea. El ciclo de desarrollo de C. burnetii comienza con la entrada de las variantes celulares pequeñas (SCV y SDC) en la célula eucariota por endocitosis (Baca & Paretsky, 1983). Una vez dentro de la célula acidifican el endosoma hasta un pH de 5,5, para luego multiplicarse por fisión binaria y comenzar a diferenciarse en las variantes celulares grandes (LCV). Transcurrido un tiempo tras la endocitosis, el fagosoma que contiene formas celulares grandes (LCV) se fusiona con los lisosomas que acidifican el medio hasta un pH de 4,5. Este pH es necesario para que la bacteria pueda activar su metabolismo (Heinzen et al., 1999) y para que las

LCV se multipliquen, proceso que está comprendido entre 1 y 2 días. Las LCV predominan durante la primera semana de infección, tiempo en el que experimentan un aumento exponencial. Al final de la primera semana de la infección se observa la fase estacionaria, donde las LCV se transforman en SCV y también se forman a modo de endosporas las SDC. Por último, se produce la liberación de las dos variantes celulares pequeñas fuera de la célula. Después de la primera replicación en los ganglios linfáticos regionales, una bacteriemia puede ocurrir. Esta bacteriemia produce que la infección se extienda a varios órganos, como hígado, bazo, médula osea, tracto reproductivo, glándulas mamarias y en hembras gestantes, a la placenta. Después de esto, aparecen lesiones granulomatosas en el hígado y la médula ósea (Maurin & Raoult, 1999;

Woldehiwet, 2004; Angelakis & Raoult, 2010). La infección crónica se establece en ciertos tejidos, incluyendo el tracto reproductivo y las glándulas mamarias.

Las hembras infectadas excretan grandes cantidades de C. burnetii durante el parto y/o restos del aborto, incluso también en heces, moco vaginal, orina y leche (Rodolakis et al.,

2007; Guatteo et al. 2007; Rodolakis, 2009). Generalmente la excreción por cualquier vía de puede durar varios meses (Berri et al., 2007) y puede ocurrir incluso en animales

43 asíntomáticos (Rousset et al., 2009). De hecho, no se observan diferencias significativas en la proporción entre cabras con y sin aborto que excretan la bacteria (Rousset et al.,

2009). La excreción en pequeños rumiantes se produce principalmente después del primer parto y/o aborto. Sin embargo, ovejas y cabras son capaces de excretar C. burnetii después del segundo y tercer parto post-infección (Berri et al., 2002; Berri et al., 2007; Rousset et al, 2009). Los patrones de excreción en animales infectados son muy variables y ciertos patrones predominantes se han identificado en el vacuno lechero (Guatteo et al., 2007), indicando la existencia de animales superexcretores, que excretan la bacteria durante varios meses e incluso a lo largo de años sucesivos (Maurin & Raoult, 1999; Guatteo et al., 2006). El conocimiento actual de las rutas de excreción en fauna silvestre se limita a la evidencia de C. burnetii en: i) secrecciones genitales de ciervo rojo, conejo de monte y jabalí en España (González-Barrio et al., 2015b,d,e,f – CAPÍTULOS II.2, III.1, III.2 y

IV), y de pequeños mamíferos como en el ratón saltador de bosque (Napaeozapus insignis), ratón ciervo de Norteamérica (Peromyscus maniculatus), topillo de espalda roja

(Myodes gapperi), ardilla roja norteamericana (Tamiasciurus hudsonicus) y ardillas voladoras (Glaucomys sabrinus y Glaucomys Volans) en Estados Unidos (Thompson et al., 2012) y en topillos campesinos en España (González-Barrio D. et al., datos sin publicar); ii) leche y/o glándula mamaria en ciervo rojo (González-Barrio et al., 2015f –

CAPÍTULO IV); iii) semen en gacela dorcas (Gazella dorcas) en un zoo en España y en ciervo en España (González-Barrio D. et al., datos sin publicar); iv) heces en cuervo de la selva (Corvus macrorhynchos) en Japón, en paloma (Columbia sp.) en Francia, en canguro gris (Macropus fuliginosus) en Australia, en perezoso de tres dedos (Bradypus trydactilus) en Guayana francesa, y en ciervo rojo y jabalí en España (To el at., 1998;

Stein et al., 1999; Banazis et al., 2010; Potter et al., 2011; Davoust et al., 2014; González-

Barrio et al., 2015a,e. CAPÍTULOS II.1 y III.2). Coxiella burnetii permanece además

44 viable e infectiva después de ser excretada en heces (Stein et al., 1999). Por lo tanto, las vías de excreción de C. burnetii en fauna silvestre son similares a las descritas en rumiantes domésticos. El vínculo entre el parto y la excreción de C. burnetii por hembras infectadas puede determinar que en especies con una época restringida de partos exista una estación predominante de excreción de C. burnetii. Esto fue observado en el brote de fiebre Q ocurrido en los Paises Bajos durante los años 2007 a 2010 en el que los casos humanos alcanzarón su punto máximo alrededor de la principal época de partos en cabras

(Roest et al., 2011a). En fauna silvestre este patrón ha sido observado en algunas especies de ungulados como el rebeco alpino (Pioz et al., 2008b) y en ciervo rojo (González-Barrio et al., 2015c – CAPÍTULO II.3). Otras especies de fauna silvestre sin una estación de partos concreta, como por ejemplo el conejo de monte, podrían excretar y transmitir C. burnetii a lo largo de casi todo el año (González-Barrio et al., 2015b – CAPÍTULO II.2).

6. Manifestaciones clínicas

Las manifestaciones clínicas de la fiebre Q en la fauna silvestre son dificiles de detectar y constituyen, en la mayoría de las ocasiones, sólo la punta del iceberg del impacto clínico esperable de la infección. En el caso particular de C. burnetii, cuyas manifestaciones clínicas conocidas en animales se limitan a fallos en la reproducción sin mortalidad materna (Clemente et al., 2008; Brom et al., 2015; García-Ispierto et al., 2014; Roest et al., 2013a), los casos son incluso más dificiles de detectar que en infecciones por otros patógenos. La mayoría de los casos de fiebre Q descritos en fauna silvestre provienen de animales en cautividad, pero el fracaso reproductivo asociado a la fiebre Q también se ha documentado en fauna silvestre en libertad. Las manifestaciones descritas de fiebre Q en fauna silvestre son: i) Fallo reproductivo - como abortos, nacidos débiles y muerte fetal - en antílope acuático (Kobus ellipsiprymnus) y en antílope sable (Hippotragus niger niger) en un zoo en Lisboa, Portugal (Clemente et al., 2008), en un rebaño de gacela dama

(Nanger dama) en cautividad en Emiratos Árabes Unidos (Lloyd et al., 2010), posiblemente en una granja de ciervo rojo en España (González-Barrio et al., 2015c –

CAPÍTULO III.2) y en una granja de búfalo de agua (Bubalus bubalis) en Italia

(Perugini et al., 2009); y ii) Placentitis, en foca común (Phoca vitulina richardsi; Lapointe et al., 1999), en león marino de Steller (Eumetopias jubatus; Kersh et al., 2010) y en oso marino ártico (Callorhinus ursinus) en Estados Unidos (Duncan et al., 2012; Myers et al.,

2013), y en ciervo rojo en Hungría (Kreizenguer et al., 2015). De los dos casos de fallo reproductivo descritos por Clemente et al., (2008) en antílope acuático y antílope sable, las madres no mostraron signos clínicos y se recuperaron bien tras el problema reproductivo. Coxiella burnetii fue confirmada por PCR en todas las muestras analizadas

(cerebro, hígado, bazo y pulmón). El estudio en fetos de búfalo de agua en la región de

Campania, el sur de Italia (Perigini et al., 2009) encontró que 14 de los 164 fetos examinados (17,5%) eran positivos a C. burnetii, lo que según los autores confirma la implicación de la fiebre Q como causa de mortalidad fetal en esta especie. Fallos reproductivos causados por fiebre Q en especies en peligro de extinción que son criados con fines de conservación pueden ser de gran preocupación, por ejemplo la reintroducción de la gacela dorcas en Senegal se vió afectada por fallos reproductivos en los que, aparte de otros patógeno, estaba implicada la fiebre Q (Teresa Abaigar, comunicación personal).

También se ha observado un brote de abortos debido a la fiebre Q en gacela dama en cautividad en los Emiratos Árabes (Lloyd et al., 2010). Estos autores describen cinco casos de aborto en la fase final de la gestación. Recientemente en España también se ha detectado la presencia de C. burnetii en semen de un macho de gacela dorcas en el zoo- aquarium de Madrid (Teresa García Seco, comunicación personal), entidad colaboradora en los programas de conservación y re-introducción de esta especie en sus zonas de origen. Este hallazgo ha alertado a los investigadores del programa de cría en cautividad,

46 junto con los problemas observados en Senegal, de la importancia de este y otros patógenos zoonóticos sobre la conservación de estas especies silvestres en grave peligro de extinción.

Mientras tanto, la infección en aves por especies similares a Coxiella, especialmente en

Psittaciformes y Piciformes en cautividad, puede tener un desenlace fatal. Infecciones por estas bacterias han sido documentadas en rosella común (Platycercus eximius), loro cacique (Deroptyus accipitrinus), chirirí (Brotogeris chiriri), perico maorí cabecirrojo

(Cyanoramphus novaezelandiae), cacatúa ninfa (Nymphicus hollandicus), Tucán

(Ramphastos toco), y loro arco iris (Trichoglossus haematodus moluccanus) (Shivaprarad et al., 2008; Woc-Colburn et a., 2008). La mayoría de las aves infectadas mostraron letargia y debilidad durante varios días antes de la muerte; signos neurológicos progresivos, presión intracraneal, hemiparesia y convulsiones.

7. Cuadro lesional

La placentitis es una de las principales lesiones observadas en los casos de fiebre Q descritos en mamíferos silvestres. Las lesiones placentarias en cabras incluyen respuesta inflamatoria severa del miometrio y la metritis es con frecuencia la única manifestación de la enfermedad en el ganado (Arricau-Bouvery and Rodolakis, 2005). En los casos en fauna silvestre se encontró una placentitis con necrosis multifocal sin evidencias de otras bacterias ni hongos (Kreizenguer et al., 2015). En un caso de aborto por fiebre Q en el

último tercio de la gestación en gacela dama no se evidenciaron lesiones post-mortem macroscópicas ni anormalidades en los fetos abortados (Lloyd et al., 2010). El examen histopatológico de los fetos reveló lesiones compatibles con sufrimiento fetal. Sí se observó en este caso una placentitis necrotizante aguda con inclusiones intracelulares correspondientes a C. burnetii. En los cotiledones de las placentas infectadas se observaron exudados y engrosamiento fibrinoso. Kersh et al. (2010) describe una

47 placentitis en león marino de Steller en la que las células trofoblásticas presentaron gran cantidad de cocobacilos fuertemente inmunorreactivos con un anticuerpo policlonal de C. burnetii. Lapointe et al. (1999) describe una placentitis causada por C. burnetii en foca común. En este caso, el citoplasma de los trofoblastos en las zonas de la membrana corioalantoidea estaba distendido por grandes formaciones esféricas. Se observó además una evidente exfoliación de trofoblastos necróticos, acumulación de material hipereosinofílico con contenido de restos nucleares en la superficie corioalantoidea y hemorragia multifocal. Las células exfoliadas contenían agregados citoplasmáticos. No se observó infiltración de células inflamatorias en la placenta.

La infección por bacterias similares a Coxiella en aves psitácidas y en tucán ocasionó una emaciación leve a moderada (Shivaprarad et al., 2008). El hígado y el bazo aparecieron gravemente agrandados, pálidos y moteados. Epicardio, endocardio, intersticio pulmonar, riñones, glándulas suprarrenales, glándulas tiroides, la lámina propia del intestino y, en algún caso, el cerebro, la bolsa de Frabicio y la médula ósea, mostraron inflamación.

Microscópicamente se describe necrosis multifocal de los hepatocitos con infiltración de células inflamatorias, entre ellos linfocitos, heterófilos, celulas plasmáticas y macrófagos, en la mayoría de las aves. Pequeños cocobacilos basófilos fueron evidentes dentro de los macrófagos de varias aves infectadas. Se observó un aumento de células del sistema fagocítico mononuclear en el bazo; algunas de estas células también contenían vacuolas con pequeños cocobacilos. Los bacilos eran evidentes por microscopía electrónica de transmisión en el hígado, los pulmones y las glándulas tiroideas. Lesiones macroscópicas, hepatomegalia y esplenomagalia se observaron en loros arco iris (Woc-Colburn et al.,

2008). El examen histopatológico en este último caso reveló microgranulomas diseminados en el hígado, el bazo y el cerebro. También se observó encefalitis vascular linfohistiocitaria y vasculitis cefálica. Mediante microscopía electrónica observaron gran

48 cantidad de macrófagos en las lesiones cerebrales, revelando organismos procariotas con forma esférica con una pared celular trilaminar en ellos.

8. Diagnóstico de la infección por Coxiella burnetii y de la fiebre Q

El diagnóstico de las infecciones y enfermedades presentes en la fauna silvestre es actualmente más complicado que en los animales domésticos y el ser humano (Arricau-

Bouvery & Rodolakis, 2005; Maurin & Raoult, 1999; Angelakis & Raoult, 2010) por diversos motivos que incluyen el acceso a casos clínicos, el acceso a muestras, la calidad de las muestras o la existencia de pruebas diagnósticas específicas. Los sistemas de vigilancia de enfermedades y patógenos de la fauna silvestre no son comparables a los existentes en el ganado, los animales domésticos y los seres humanos, incluso en países desarrollados donde hay una percepción más positiva de la salud de la fauna silvestre como parte integral de la ‘Salud Global’ (Kuiken et al., 2011). Esto es, en parte, causado por una percepción errónea de la salud de la fauna silvestre como un asunto de poca relevancia y, por lo tanto, en el que no merece la pena invertir esfuerzos. Diversos factores contribuyen a esa percepción errónea de la salud de la fauna silvestre, especialmente la escasez de conocimientos sobre la situación de los patógenos en la fauna silvestre. Una restricción para el estudio y diagnóstico de la infección por C. burnetii en la fauna silvestre es el acceso a muestras. Los programas de vigilancia pasiva de la fauna silvestre, cuando existen y funcionan, recolectan muestras de animales encontrados muertos o enfermos (MAGRAMA). Habitualmente, los cadéveres de animales muertos se localizan bastante tiempo después de la muerte del animal, por lo que en la mayor parte de los casos las muestras son de mala calidad para el diagnóstico. Por otro lado, los programas de vigilancia activa de la fauna silvestre, cuando existen, suelen proporcionar muestras de animales recolectadas de forma oportunista y sólo representativas de las poblaciones a escalas geográficas demasiado grandes como para ser representativas de la situación real.

Muchos patógenos causantes de enfermedad en animales domésticos y humanos han evolucionado con la fauna silvestre y son de baja patogenicidad en estas especies. Por ejemplo, las cepas del virus de la enfermedad de Aujeszky (ADV) que están presentes en el cerdo doméstico presentan mayor virulencia que las cepas de ADV circulantes en las poblaciones de jabalí (Muller et al., 2010, 2011). Por lo tanto, mientras que las manifestaciones clínicas del ADV son más propensas a ser observadas en cerdos domésticos, estas rara vez se observan en jabalí. Infecciones por patógenos más virulentos en el jabalí como el virus de la peste porcina africana (Tauscher et al., 2015), son más evidentes por la mortalidad evidente que conllevan. Algo similar a ADV parece ocurrir con C. burnetii debido al bajo porcentaje de hembras infectadas que experimentan fallo reproductivo (Arricau-Bouvery & Rodolakis, 2005).

Por otro lado, la detección de casos clínicos y por lo tanto, la disponibilidad de las muestras necesarias para el diagnóstico, es más elevada en grupos de animales controlados por el ser humano. Como ejemplo, los casos de fiebre Q en colecciones zoológicas son detectados por los cuidadores que revisan el estado de los animales con frecuencia diaria y pueden detectar síntomas de malestar en los animales o fallos reproductivos, y con ello acceder rápidamente a muestras en buen estado de conservación.

Por el contrario, y tomando como ejemplo una población de ciervos en producción extensiva en granja, mucho más vigilada que cualquier población silvestre de ciervos, detectar fallo reproductivo por fiebre Q y acceder a las muestras (en buena o mala calidad) es una árdua tarea y suelen pasar desapercibidas (González-Barrio et al., 2015c –

CAPÍTULO III.2). En fauna silvestre en libertad esto resulta aún más complicado, ya que los restos del parto, de animales mortinatos o de anejos fetales son encontrados muy rara vez y de forma casual (Ruiz-Fons et al. 2006).

Finalmente, aunque algunas técnicas diagnósticas son igualmente válidas para humanos, animales domésticos y silvestres – aislamiento, PCR, immunohistoquímica, etc. – otras como las técnicas de diagnóstico serológico, son sólo válidas para las especies para las que están diseñadas debido a la variabilidad en la estructura de los anticuerpos entre especies. A continuación se recogen las diferentes técnicas disponibles para el diagnóstico de la infección por C. burnetii y su aplicación en animales silvestres:

Métodos directos

Aislamiento. El aislamiento de C. burnetii se puede realizar mediante modelos animales in vivo, cultivos celulares o huevos de aves embrionados, siempre que la carga de C. burnetii en la muestra y la contaminación por otras bacterias lo permitan (Ho et al., 1995).

Los modelos animales in vivo han sido muy utilizados, aunque su uso como modelo de diagnóstico no es recomendable en la actualidad. La técnica de asilamiento de C. burnetii en cultivos celulares está basada en la inoculación de la muestra problema sobre células en cultivo sensibles a la infección por C. burnetii como células Vero o células de fibroblastos pulmonares de embrión humano - células HEL. El efecto citopático que la infección por C. burnetii provoca sobre las células de cultivo conduce a la formación de vacuolas, señal de que la infección ha tenido lugar. La principal ventaja de esta técnica es su rápidez para el aislamieto de C. burnetii (Spyridaki et al., 2002). La principal desventaja es la necesidad de instalaciones de alto nivel de seguridad biológica – nivel 3 o superior – para el aislamiento de la bacteria. Esta técnica ha sido utilizada tanto en muestras procendentes de humanos (Raoult et al., 1991), como de animales domésticos y silvestres o garrapatas (Psaroulaki et al., 2006; Spyridaki et al., 2002). La principal dificultad en el uso de esta técnica con muestras de animales silvestres, sobre todo cuando no se trate de casos clínicos, es que la cantidad de bacterias de C. burnetii en la muestra no sea la adecuada (CAPÍTULOS II.4 y II.5) y que, por lo tanto, se incremente el

51 porcentaje de resultados falsos negativos. En los últimos años se ha descrito la utilidad de un medio axénico para el aislamiento de C. burnetii en condiciones microaerófilas

(Omsland et al., 2009; Lagier et al., 2015).

El asilamiento en huevos embrionados es una técnica con una elevada sensibilidad para la detección de C. burnetii y que se puede utilizar con una amplia gama de muestras provenientes de heces, hisopos y tejidos, entre otros. La especificidad no estan alta en comparación con su sensibilidad ya que otros agentes presentes en la muestras pueden crecer también en el huevo embrionado, por este motivo la muestra a analizar debe presentar un nivel muy bajo de contaminación por otras bacterias. Es una técnica muy laboriosa y no se recomienda como técnica de rutina de diagnóstico (Klee et al., 2006;

Maurin & Raoult, 1999). Con muestras contaminadas como placentas, descargas vaginales, heces o leche, es necesario un paso previo de inoculación en animales de laboratorio, como ratones o cobayas, y luego realizar el cultivo a partir del bazo de estos animales (Scott et al., 1987).

Tinción. La detección de C. burnetii mediante esta técnica se suele realizar a partir de improntas de placentas de animales abortados. Esta técnica podría ser útil para animales silvestre que se encuentran en cautividad y de fácil acceso a muestras, pero no sería de utilidad para animales en libertad por la dificultad de acceso a muestras con altas cargas de C. burnetii. Las tinciones de uso más habituales son Stamp, Giménez, Machiavello,

Giemsa o Ziehl-Neelsen modificado (Giménez, 1965; Sanford et al., 1994). Estas tinciones permiten visualizar al microscopio de manera rápida y sencilla las formas compatibles con C. burnetii en exudados o en áreas de inflamación de la placenta. Este método carece de buena especificidad y sensibilidad ya que C. burnetii puede confundirse en ocasionaes con Chlamydophila abortus o con bacterias del género Brucella. Por este

52 motivo es recomendable complementar el diagnóstico con otras técnicas de detección más específicas de C. burnetii como la PCR.

Histología e inmunohistoquímica. Al igual que con la tinción, la detección de bacterias compatibles con C. burnetii por técnicas histológicas en tejidos puede ser indicativo de una potencial infección. Sin embargo, esta técnica necesita de técnicas confirmatorias posteriores. Las lesiones ocasionadas por C. burnetii no son patognomónicas, aunqeu la detección de placentitis necrótica supurativ, con infiltado inflamatorio (Lapointe et al.,

1999; Kersh et al., 2010) e hipertrofia de trofoblastos en los que se observan bacterias intracelulares podría ser indicativo de infección por esta bacteria.

La técnica de inmunohistoquímica se realiza a partir de tejidos includos en parafina o en frotis fijados con acetona (Raoult et al., 1994). Se basa en la realización de la técnica de inmunofluorescencia indirecta o la técnica de inmunoperoxidasa usando anticuerpos policlonales frente a C. burnetii que posteriormente son revelados. La inmunohistoquímica permite localizar con precisión los tejidos y células donde se localiza

C. burnetii. El inconveniente es que es una técnica cara, laboriosa y además no es lo suficientemente sensible como para detectar el antígeno en otros órganos distintos a la placenta (Sánchez et al., 2006) donde la carga bacteriana es más baja. En fauna silvestre su utilidad estaría restringida a casos clínicos en los que se puedan obtener muestras de buena calidad, a estudios de patogenia en infecciones experimentales y al estudio de tejidos obtenidos de animales infectados en sistemas de vigilancia activa. Sin embargo, otras técnicas complementarias serían necesarias para mejorar la sensibilidad y la especificidad del diagnóstico.

Técnicas moleculares. El desarrollo de técnicas moleculares ha supuesto un gran salto en el diagnóstico de patógenos, tanto en la medicina humana como veterinaria, permitiendo identificar con altos niveles de especificidad y sensibilidad los agentes patógenos

53 presentes en las muestras clínicas y ambientales. La técnica molecular más utilizada es la reacción en cadena de la polimerasa (PCR), basada en la amplificación de fragmentos de

ácido desoxirribonucleico (ADN) que permite generar un número exponencial de copias de un fragmento específico del patógeno (Sambrook et al., 1989). La PCR es actualmente la técnica más adecuada para detectar C. burnetii en cualquier tipo de muestra biológica

(Berri et al., 2000), aunque se debe tener en cuenta la presencia de posibles inhibidores de la reacción que interfieren en el proceso de amplificación (Capuano et al., 2004;

Lorenz et al., 1998). Sin embargo, esto se puede evitar desarrollando protocolos óptimos para la extracción de ADN. Con la base de la PCR convencional, se han desarrollado

PCRs más sensibles para la detección de C. burnetii. La PCR anidada consiste en amplificar una secuencia pequeña incluida dentro de una secuencia mayor previamente amplificada (Parisi et al., 2006; Spyridaki et al., 2002; Zhang et al., 1998). Su principal desventaja es que se duplica la probabilidad de contaminación ya que hay que manipular el producto amplificado obtenido en la primera amplificación (Parisi et al., 2006). Otra variante es la “PCR touchdown”, técnica que se emplea cuando se desconoce la secuencia exacta de los extremos de la secuencia a amplificar, de modo que se asume que puede existir alguna base desapareada en el alineamiento cebador-secuencia. Su finalidad es reducir el fondo no específico bajando gradualmente la temperatura de hibridación a lo largo del progreso de la PCR. Esta PCR es una de las más utilizadas para la identificación de C. burnetii (Berri et al., 2000; Willems et al., 1994). La PCR a tiempo real es una técnica rápida que reduce el riesgo de contaminación y tiene la capacidad de permitir la cuantificación o semi-cuantificación de la concentración de ADN diana (de Bruin et al.,

2011; Jones et al., 2011; Tilburg et al., 2010). Sus principales ventajas son la elevada especificidad y sensibilidad, la obtención de resultados en tiempo real (en un espacio reducido de tiempo) y la posibilidad de automatizar el proceso para el análisis de grandes

54 cantidades de muestras. La mayoría de las PCR a tiempo real están basadas en sondas específicas que utilizan una sonda unida a dos fluorocromos que hibrida en la zona intermedia entre el cebador directo (forward) y el inverso (reverse). Cuando la sonda está intacta, presenta una transferencia energética de fluorescencia por resonancia (FRET).

Dicha FRET no se produce cuando la sonda está dañada y los dos fluorocromos están distantes, producto de la actividad 5'-3' exonucleasa de la ADN polimerasa. Esto permite monitorizar el cambio del patrón de fluorescencia y deducir el nivel de amplificación del gen. Para la amplificación y detección de C. burnetii se han empleado diversas dianas o genes, como com1, que codifica una proteína de 27kDa de la membrana externa (Lockhart et al., 2011), el gen de la superóxido dismutasa (sodB) (Stein y Raoult, 1992), el operón de choque térmico que codifica dos proteínas de choque térmico, hspA y hspB (Fournier

& Raoult, 2003), el gen de la isocitrato deshidrogenasa (icd) (Klee et al., 2006; Nguyen et al., 1999), la proteína potenciadora de la infectividad del macrófago (cbmip) (Klee et al., 2006; Zhang et al., 1998), la secuencia de inserción en multicopias IS1111 del gen de la transposasa y sus variantes (IS1111a) (Berri et al., 2000; Lorenz et al., 1998; Willems et al., 1994, Tilburg et al., 2010). Se han descrito además, otras técnicas moleculares que ofrecen el aumento de sensibilidad, reducción del equipamiento y del tiempo de obtención de resultados, además de poner analizar un número elevado de secuencias génicas (Beare et al., 2006; Jado et al., 2012).

Una vez detectada la bacteria en muestras biológicas, podemos caracterizar el tipo de cepa presente en dichas muestras biológicas mediante técnicas de caracterización molecular.

La caracterización molecular de C. burnetii mediante genotipado es un instrumento para el reconocimiento de la diversidad de cepas circulantes. Esta es una herramienta indispensable para las investigaciones epidemiológicas de brotes de fiebre Q y para su vigilancia y control. Durante los últimos años varias técnicas de genotipado de C. burnetii

55 han sido publicadas (Sidi-Boumedine & Rousset, 2011; Massung et al., 2012). Métodos como el análisis de la longitud de los polimorfismos mediantes encimas de restricción

(PCR-RFLP) utilizando los genes icd y com 1 (Andoh et al., 2004; Spyridaki et al., 1998;

Stein & Raoult, 1992; Nguyen et al., 1999) y el método de la técnica de campo pulsado

(PFGE), en el que se requiere de una buena separación de los diferentes fragmentos

(Heinzen et al., 1990; Jäger et al., 1998), han sido usados para reconocer los diferentes grupos de aislados de C. burnetii. La diferenciación de cepas puede incluso conseguirse por secuencias basadas en la determinación de genes de codificación como Com1 y Mucz, y otros métodos comparando el genoma completo usando métodos basados en microarrays descritos en diferentes estudios de plásmidos (Jäger et al., 2002; Samuel et al., 1985; Hendrix et al., 1991; Jäger et al., 1998; Thiele et al., 1993; Nguyen et al., 1998).

Además, fragmentos de restricción infrecuentes-PCR (IRS-PCR), PCR basada en la frecuencia de inserción IS1111 y espectrometría de masas acoplada a cromatografía líquida (LC-MS/MS) han sido desarrollados para tipar los asilados de C. burnetii

(Arricau-Bouvery et al., 2006; Denison et al., 2007; Hernychova et al., 2008). Todos estos métodos necesitan un cultivo previo de la bacteria en células o huevos embrionados en condiciones de nivel 3 de bioseguridad antes de poder anilizar las muestras, condiciones que sólo existen en determinados laboratorios. Sin embargo, existen otros métodos de genotipado en los se pueden usar las muestras directamente sin tener que desarrollar un cultivo previo, en particular MST (Multispacer Sequence Typing), técnica basada en la secuanciación de diferentes regiones intergénicas que permite separar los aislados según los diferentes tipos de secuencias que muestran en las diferentes regiones. La razón para secuenciar estas regiones es que son potencialmente variables y están sujetas a una menor presión que los genes adyacentes. Los genotipos descritos mediante este método se pueden agrupar en tres grandes grupos (Glazunova et al., 2005). Otro método dentro de

56 este grupo es MLVA (Multiple Loci Variable number of tandem repeats Analysis) que permite la detección de los polimorfismos en las secuencias repetidas en tándem en el

ADN. Se han descrito un total de 17 marcadores diferentes para las repeticiones de minisatélites y microsatélites (Arricau-Bouvery et al., 2006). Otra técnica en la que también se pueden utilizar las muestras clínicas sin necesidad de preparaciones en condiciones de cultivo celular es PCR-RLB (Multiplex PCR and Reverse Line Blot

Hybridization) método menos habitual pero que permite agrupar más los genotipos, en este caso grupos genómicos, que MLVA, y con capacidad para poder comparar mayor número de cepas (Beare et al., 2006; Jado et al., 2012). Actualmente, se están llevando a cabo estudios en los que se ha puesto a punto la técnica de tipado conocida como SNP

(Single Nucleotide Polymorphism) (Hermans et al., 2011; Hornstra et al., 2011;

Huijsmans et al., 2011), considerada como una técnica rápida, sensible, fácil de llevar a cabo y que consiste en la detección de la variación de una sola base en una determinada secuencia de ADN. Esta técnica se ha usado para tipar directamente C. burnetii en muestras animales y ha sido aplicada recientemente para establecer los diferentes genotipos implicados en el brote de fiebre Q humana en Holanda (Huijsmans et al., 2011).

Además, con esta técnica se ha descrito por primera vez la detección en leche de ADN procedente de la vacuna inactivada en fase I en animales recientemente vacunados

(Hermans et al., 2011). A pesar de que tanto las técnicas MST, MLVA, PCR-RLB y SNP tienen un alto poder discriminatorio, la técnica MLVA y SNP son superiores (Arricau-

Bouvery et al., 2006, Massung et al., 2012). Además, la técnica MLVA es menos laboriosa y no necesita de secuenciación posterior. Con estas técnicas se puede analizar el ADN que ha sido extraído a partir de muestras recogidas de animales, como moco vaginal, leche, heces y cualquier tejido infectado, sin la necesidad de aislar la cepa previamente. De esta manera, se puede analizar la circulación de C. burnetii en las

57 poblaciones de animales domésticos, fauna silvestre y humanos, permitiendo elucidar cuales son las cepas predominantes en un área y especie determinada.

Existe poca información de las cepas presentes en la fauna silvestre, lo que puede ser debido a: i) La dificultad en la detección de animales de especies silvestres con infección activa por C. burnetii; y ii) Que la carga bacteriana encontrada en fauna silvestre infectada por C. burnetii no es suficientemente elevada como para poder realizar la caracterización, ya que algunas técnicas necesitan una cantidad mínima de bacterias para poder caracterizar por completo la cepa. Por estos motivos los estudios en los que se han caracterizado cepas de C. burnetii en fauna silvestre son muy reducidos. Entre ellos dos estudios fueron realizados tras brotes de fiebre Q en humanos, uno en Holanda (Rijks et al., 2011) y otro en Guayana Francesa (Davoust et al., 2014). En el primer caso el corzo fue objeto de estudio, analizando corzos de diferentes regiones en los Países Bajos, en los que se encontró un 23% de infección por C. burnetii; sin embargo a la hora de caracterizar las cepas presentes en estos animales no se obtuvieron genotipos completos, probablemente debido a la baja carga bacteriana en las muestras y a la utilización de

MLVA. En otro estudio (Davoust et al., 2014) la técnica utilizada fue MST, detectando

C. burnetii en heces y garrapatas de perezoso de tres dedos. En este caso el genotipo encontrado tanto en las garrapatas como en las heces del animal coincidía con el genotipo detectado en personas y en animales domésticos, el único genotipo circulante. Estudios recientes en fauna silvestre utilizan tanto MLVA como PCR-RLB para estudiar las cepas de C. burnetii circulantes en humanos y animales (Jado et al., 2012; Cumbassa et al.,

2015; CAPÍTUOS II.4 y II.5).

Métodos indirectos

Técnicas serológicas. Entre las técnicas serológicas más usadas para el diagnóstico de la fiebre Q tenemos; i) La inmunofluorescencia indirecta (IFA); ii) La fijación del

58 complemento (CFT); y iii) El ensayo de inmunoabsorbancia enzimática (ELISA)

(Herremans et al., 2013). Tanto ELISA como CFT son ampliamnete utilizados en el diagnóstico veterinario de exposición a C. burnetii, mientras que IFA es principlamente utilizado en medicina humana. La sensibilidad difiere entre las técnicas, siendo IFA y

CFT - especialmente esta última - menos sensibles que el ELISA (Astobiza et al., 2007;

Rousset et al., 2007; Kittelberger et al., 2009). La utilización de ELISA es más común en estudios epidemiológicos, ya que sus protocolos están estandarizados entre laboratorios.

Actualmente existen varios ELISA comerciales para detectar anticuerpos frente a C. burnetii en rumiantes domésticos, y que con ciertas modificaciones tambíen pueden utilizarse en fauna silvestre (González-Barrio et al., 2015a,b – CAPÍTULOS II.1 y II.2).

Se ha demostrado que esta técnica puede mostrar diferentes sensibilidades (Kittelberger et al., 2009). Así, el ELISA preparado con antígeno de aislados de rumiantes tiene una mayor sensibilidad que los que se preparan con antígenos procedentes de aislados de garrapatas (Rodolakis 2006; Rodolakis et al., 2007). En humanos el diagnóstico serológico de la fiebre Q aguda o crónica se ha basado principalmente en la detección de anticuerpos de fase I o de fase II mediante IFA, y en la relación entre ambos, ya que esta varía en función del curso de la infección. En ELISAs comerciales para uso veterinario no se se hace habitualmente esta diferencia ya que se tapizan los pocillos con antígenos de fase I y II al tiempo. Sin embargo, sí existen ELISAs comerciales específicos para la detección de los anticuerpos específicos para cada tipo de antígeno de C. burnetii.

Algunas investigaciones han analizado por separado la presencia de los dos antígenos

(Böttcher et al., 2011; Cooper et al., 2011). La interpretación de los resultados de ELISA a nivel individual puede resultar dificil ya que hay animales que pueden permanecer seropostivos durante años tras una infección, eliminando o no la bacteria, pero también algunos animales infectados no llegan a seroconvertir (Arricau-Bouvery et al., 2003;

Berri et al., 2001). También se plantea otro problema en la detección de anticuerpos por

ELISA, ya que el ELISA no es capaz de diferenciar anticuerpos vacunales de anticuerpos debidos a infección. De este modo la manera más adecuada para hacer un diagnóstico fiable sería combinar varias técnicas, en este caso ELISA y PCR (Arricau-Bouvery y

Rodolakis, 2005).

9. Prevención y control

El elevado impacto de la fiebre Q sobre la salud pública y la sanidad animal ha propiciado el desarrollo de herramientas para la prevención y control de la enfermedad. En humanos, el estudio de terapias antibióticas preventivas para el control de la infección por C. burnetii ha sido ampliamente desarrollado. A la par, se sigue investigando sobre el desarrollo de vacunas y protocolos de vacunación eficaces en humanos que prevengan la infección por C. burnetii. Como fuente primordial de infección para el ser humano y por los problemas que la infección causa en la producción animal, también se han desarrollado numerosas herramientas preventivas y de control frente a C. burnetii en animales domésticos, especialmente en especies de rumiantes.

En el ganado las medidas de control se basan en la profilaxis, como mejoras en la higiene, mejoras en los sistemas de bioseguridad o en el desarrollo de vacunas y protocolos de vacunación como medida preventiva. Como medidas de control de la infección se han ensayado algunos tratamientos. Existen algunos tratamientos con antibióticos

(especialmente tetraciclinas) que se pueden utilizar en animales domésticos, si bien su efectividad es cuestionable (Astobiza et al., 2011b). La oxitetraciclina usada como tratamiento en los rebaños infectados de ovejas no parece tener un efecto en la reducción de la excreción ni en el número de animales excretores (Astobiza et al., 2010). Sin embargo, la vacunación con vacunas inactivadas de fase I después de aplicar el tratamiento con oxitetraciclina reduce el porcentaje de excretores a largo plazo (Astobiza

60 et al., 2013). La vacunación parece ser una de las medidas más eficaces para controlar la enfermedad en los rumiantes domésticos (Arricau-bouvery et al., 2005). Así, las vacunas preparadas a partir de la fase antigénica I han demostrado ser más eficaces que las preparadas con la fase antigénica II (Arricau-bouvery et al., 2005), y se ha demostrado su eficacia en rebaños de vacas, ovejas y cabras infectados como protectora de la infección en individuos no infectados (Guatteo et al, 2008; Hogerwerf et al, 2011; Rousset et al,

2009b.) Sin embargo, aunque la vacunación reduce el riesgo de infección en los animales no infectados (Guatteo et al., 2008), en rebaños altamente infectados la vacunación no tiene un efecto significativo inmediato y, aunque existe una disminución sucesiva de la excreción y de la carga bacteriana - también del número de abortos, la seroconversión de animales no vacunados indica la presencia de infección activa en el rebaño (Astobiza et al., 2011b). Además no se observan diferencias significativas entre lotes vacunados y no vacunados a nivel de excreción, sugeriendo que en rebaños con alto nivel de infección la vacuna a corto plazo no es efectiva (Astobiza et al., 2011b).

Hasta la fecha no se ha analizado ningún protocolo de prevención y control de la infección por C. burnetii en la fauna silvestre salvo el estudio llevado a cabo en esta Tesis Doctoral

(CAPÍTULO V). Sin embargo, el desarrollo de estrategias de control de patógenos en la fauna silvestre es una disciplina emergente. La mayor parte de las actuaciones de control y erradicación de enfermedades en fauna silvestre se ha realizado sobre enfermedades con impacto en la conservación de especies en peligro o sobre enfermedades compartidas con los animales domésticos y/o el hombre en las que el papel de la fauna se ha demostrado como relevante (Gortázar et al., 2011). El control de las enfermedades compartidas por la fauna silvestre requiere el desarrollo de estrategias para reducir la transmisión del patógeno individuos, bien sea controlando la proporción de individuos infectados en la población (eliminación no selectiva, testaje y eliminación, tratamientos,

61 etc.), bien sea protegiendo a los individuos susceptibles frente a la infección (vacunas, medidas de bioseguridad, higiene, etc.) o ambas de forma integral. El control de las enfermedades en la fauna silvestre a menudo consiste en la intervención en los ecosistemas naturales, lo que resulta en numerosas ocasiones controvertido (Artois et al.,

2011). Este control de las enfermedades en fauna silvestre ha sido ampliamente descrito en la literatura y los diferentes métodos estudiados son: i) Acciones preventivas, entre las que destacan el control del movimiento de animales (Gilbert et al., 2005; Cartersen et al.,

2011) así como el control sanitario previamente al movimiento para evitar la introducción o re-introducción de patógenos a través de animales infectados o cuando animales silvestres mantenidos en cautividad son liberados. Otro método de prevención activa es la utilización de barreras que dificulten el contacto entre animales sanos y animales infectados (medidas de bioseguridad), como los cercados, que son útiles contra patógenos de transmisión directa como la fiebre aftosa (Sutmoller et al., 2002; Scheider et al., 2012) o la tuberculosis bovina (Judge et al., 2011) al impedir el movimiento de los animales

(Owens & Owens, 1980). En el caso de C. burnetii que se puede transmitir a través de aerosoles las barreras no deben ser muy eficaces; la bacteria puede ser vehiculada a varios kilometros de distancia por el viento (Nusinovici et al., 2015); ii) El control poblacional es otra medida eficaz para prevenir la transmisión de patógenos de animales infectados a sanos al reducir la prevalencia (Boadella et al., 2012a). Los métodos de control poblacional sin una diana clara en animales infectados pueden ser controvertidos, pero también se han desarrollado protocolos de testaje y eliminación selectiva de animales infectados que han resultado ser eficaces (Shury et al., 2015); y iii) La vacunación en fauna silvestre también puede ser una herramienta útil y complementaria para reducir la prevalencia de la infección en poblaciones de animales silvestres (Beltrán-Beck et al.,

2012). Este método ha sido eficaz en el control de la rabia en el zorro rojo (Vulpes vulpes) en el este de Europa (Monnerot et al., 2015).

Tabla 1, 2, 3, 4 y 5. Especies de mamíferos, aves, anfibios, reptiles, peces, artrópodos y especies silvestres en cautividad en los que se ha detectado tanto anticuerpos frente a Coxiella burnetii como ADN.

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference France 5/61 (8.2) CFT Baradel et al., 1988 Alpine ibex Capra ibex http://maps.iucnredlist.org/map.html?id=42397 0/10 (0.0) PCR Marreros et al., 2011 Switzerland 10/551 (1.8) ELISA Marreros et al., 2011 Aoudad Ammotragus lervia http://maps.iucnredlist.org/map.html?id=1151 Poland 0/1 (0.0) ELISA Rypula et al., 2011 Bighorn sheep Ovis canadensis http://maps.iucnredlist.org/map.html?id=15735 USA 27/268 (10.0) CFT Deforge et al., 2006 8/135 (5.9) CFT Baradel et al., 1988 n.a. (0.0-50.0) CFT Jourdain et al., 2005 Alpine chamois Rupicapra rupicapra http://maps.iucnredlist.org/map.html?id=39255 France 0/125 (0.0) CFT Pioz et al., 2008a n.a. (0.0-12.0) CFT Pioz et al., 2008b Chinkara Gazella bennettii http://maps.iucnredlist.org/map.html?id=8978 India n.a. ISO Yadav and Sethi, 1980 Common wildebeest Connochaetes taurinus http://maps.iucnredlist.org/map.html?id=5229 Poland 0/1 (0.0) ELISA Rypula et al., 2011 Thinhorn sheep Ovis dalli http://maps.iucnredlist.org/map.html?id=39250 USA 5/15 (33.3) CFT Zarnke et al., 1983 Artiodactyla Bovidae 0/122 (0.0) SERO Salwa et al., 2007 0/40 (0.0) ELISA Rypula et al., 2011 European bison Bison bonasus http://maps.iucnredlist.org/map.html?id=2814 Poland 36/47 (76.6) SERO Skarek et al., 1994 7/60 (11.6) CFT/MAT Kita et al., 1991 Spanish ibex Capra pyrenaica http://maps.iucnredlist.org/map.html?id=3798 Spain 7/52 (13.4) ELISA Santiago-Moreno et al., 2011 Japanese serow Capricornis crispus http://maps.iucnredlist.org/map.html?id=3811 Japan 0/117 (0.0) ELISA Ejercito et al., 1993 23/74 (31.1) PCR Psaroulaki et al., 2014 Cyprus 23/77 (29.8) PCR Ioannou et al., 2011 Mouflon Ovis orientalis http://maps.iucnredlist.org/map.html?id=15739 2/2 (100.0) MAT Hubalez et al., 1993 Czech Rep. Spain 4/101 (3.9) ELISA Lopez-Olvera et al., 2009 Muskox Ovibos moschatus http://maps.iucnredlist.org/map.html?id=29684 Canada 5/17 (29.4) SERO Seguin et al., 2008 Boselaphus Nilgai http://maps.iucnredlist.org/map.html?id=2893 Poland 0/1 (0.0) ELISA Rypula et al., 2011 tragocamelus

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference Sitatunga Tragelaphus spekii http://maps.iucnredlist.org/map.html?id=22050 Poland 0/1 (0.0) ELISA Rypula et al., 2011 Bovidae Yak Bos mutus http://maps.iucnredlist.org/map.html?id=2892 China 75/552 (13.6) ELISA Yin et al., 2015 Moose Alces americanus http://maps.iucnredlist.org/map.html?id=818 Canada n.a. (16.5) IFA Marrie et al., 1993 Odocoileus hemionus Black-tailed deer http://maps.iucnredlist.org/map.html?id=42393 USA 5/143 (3.5) ELISA Chomel et al., 1994 colombianus California mule deer O. h. californicus http://maps.iucnredlist.org/map.html?id=42393 USA 1/26 (3.8) ELISA Chomel et al., 1994 Axis deer Axis axis http://maps.iucnredlist.org/map.html?id=41783 Poland 0/2 (0.0) ELISA Rypula et al., 2011 Eurasian elk Alces alces http://maps.iucnredlist.org/map.html?id=41782 Sweden 0/99 (0.0) CFT Ohlson et al., 2014 Czech Rep. 2/4 (50.0) MAT Hubalek et al., 1993 Hungary 0/22 (0.0) PCR Kreizinger et al., 2015 Fallow deer Dama dama http://maps.iucnredlist.org/map.html?id=42188 Italy 3/43 (6.9) CFT Giovannini et al., 1988 Spain 0/13 (0.0) IFA Ruiz-Fons et al., 2008 Artiodactyla 42/61 (68.8) ELISA Ejercito et al., 1993 Cervidae Sika deer Cervus nippon http://maps.iucnredlist.org/map.html?id=41788 Japan 0/5 (0.0) IFA Neagary et al., 1998 n.a (22.0) CFT Enright et al., 1971 Wapiti Cervus canadensis http://maps.iucnredlist.org/map.html?id=41785 USA n.a. SERO McQuiston et al., 2002 Czech Rep. 6/24 (25.0) MAT Hubalek et al., 1993 Slovakia 1/3 (33.3) PCR Smetanova et al., 2006 0/28 (0.0) PCR Astobiza et al., 2011 Red deer Cervus elaphus http://maps.iucnredlist.org/map.html?id=41785 1/2 (50.0) PCR Ruiz-Fons et al., 2008 Spain 2/36 (5.5) IFA Ruiz-Fons et al., 2008 27/460 (5.8) PCR González-Barrio et al., 2015c 41/1151 (3.5) ELISA González-Barrio et al., 2015c

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference France 1/54 (1.8) CFT Baradel et al., 1988 Red deer Cervus elaphus http://maps.iucnredlist.org/map.html?id=41785 Hungary 2/36 (5.5) PCR Kreizinger et al., 2015 Reindeer Rangifer tarandus http://maps.iucnredlist.org/map.html?id=29742 Poland 0/2 (0.0) ELISA Rypula et al., 2011 Rocky mountain mule O. h. hemionus http://maps.iucnredlist.org/map.html?id=42393 USA 6/60 (10.0) ELISA Chomel et al., 1994 deer 25/222 (11.2) ELISA Candela et al., 2014

France 26/697 (3.7) CFT Baradel et al., 1988

3/175 (1.7) CFT Blancou et al., 1983

Hungary 0/33 (0.0) PCR Kreizinger et al., 2015

Netherl. 18/79 (23.0) PCR Rijks et al., 2011

Roe deer Capreolus capreolus http://maps.iucnredlist.org/map.html?id=42395 Poland 0/20 (0.0) ELISA Rypula et al., 2011 Cervidae Czech Rep. 2/33 (6.0) MAT Hubalek et al., 1993

Slovakia 0/2 (0.0) PCR Smetanova et al., 2006 Artiodactyla 0/6 (0.0) PCR Ruiz-Fons et al., 2008

Spain 4/78 (5.1) PCR Astobiza et al., 2011

6/39 (15.4) IFA Ruiz-Fons et al., 2008

Southerm mule deer O. h. fuliginatus http://maps.iucnredlist.org/map.html?id=42393 USA 0/47 (0.0) ELISA Chomel et al., 1994

Canada n.a. (1.5) IFA Marrie et al., 1993 White-tailed deer Odocoileus virginianus http://maps.iucnredlist.org/map.html?id=42393 USA 112/624 (17.9) IFA Kirchgessner et al., 2012

13/196 (6.6) PCR Shin et al., 2014 Water deer Hydropotes inermis http://maps.iucnredlist.org/map.html?id=10329 Korea 18/196 (9.2) ELISA Shin et al., 2014

Czech Rep. 2/32 (6.2) MAT Hubalek et al., 1993

France 0/209 (0.0) CFT Baradel et al., 1988 Suidae Eurasian wild boar Sus scrofa http://maps.iucnredlist.org/map.html?id=41775 Germany n.a. (2.2-8.1) SERO Hening et al., 2015

Germany 6/220 (2.7) SERO Hartung, 2001

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference Italy 0/20 (0.0) CFT Giovannini et al., 1988 Japan 0/30 (0.0) ELISA Ejercito et al., 1993 Slovakia 0/18 (0.0) PCR Smetanova et al., 2006 Eurasian wild boar 1/1 (100.0) PCR Jado et al., 2012 Artiodactyla Suidae Sus scrofa http://maps.iucnredlist.org/map.html?id=41775 1/125 (0.8) ELISA González-Barrio et al., 2015d Spain 4/112 (3.6) PCR González-Barrio et al., 2015d 4/93 (4.3) PCR Astobiza et al., 2011 67/135 (50.0) SERO Clark et al., 1983 Wild swine USA n.a. CFT Sidwell et al., 1964 African wild dog Lycaon pictus http://maps.iucnredlist.org/map.html?id=12436 S. Africa 8/29 (27.6) SERO Van Heerden et al., 1995

n.a. (63.0) MAT Willeberg et al., 1980

Coyote Canis latrans http://maps.iucnredlist.org/map.html?id=3745 USA n.a. (78.0) CFT Enright et al., 1971

n.a. SERO McQuiston et al., 2002

Darwin´s fox Pseudalopex fulvipes http://maps.iucnredlist.org/map.html?id=41586 Cyprus 0/30 (0.0) PCR Cabello et al., 2013

Nyctereutes 0/30 (0.0) IFA Neagary et al., 1998 Japanese racoon dog http://maps.iucnredlist.org/map.html?id=14925 Japan procyonoides viverrinus 0/37 (0.0) ELISA Ejercito et al., 1993

Canidae Czech Rep. 1/1 (100.0) MAT Rehacek et al., 1977 Carnivora Cyprus 9/32 (28.1) PCR Psaroulaki et al., 2014

Portugal 0/4 (0.0) PCR Cumbassá et al., 2015

Spain 0/61 (0.0) PCR Astobiza et al., 2011 Red fox Vulpes vulpes http://maps.iucnredlist.org/map.html?id=23062 UK 32/120 (26.6) ELISA Meredith et al., 2014

n.a. (55.0) CFT Enright et al., 1971

USA n.a. (63.0) MAT Willeberg et al., 1980

n.a. SERO McQuiston et al., 2002

Felidae Bobcat Lynx rufus http://maps.iucnredlist.org/map.html?id=12521 USA n.a. CFT Enright et al., 1971

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference USA n.a. CFT Enright et al., 1971 Feral cat Felis silvestris catus India n.a. (0.0) CAT Yadav and Sethi, 1980 Felidae Tiger Panthera tigris http://maps.iucnredlist.org/map.html?id=15955 India n.a. (0.0) CAT Yadav and Sethi, 1980 Wild cat Felis silvestris http://maps.iucnredlist.org/map.html?id=60354712 Spain 0/6 (0.0) PCR Astobiza et al., 2011 Egyptian mongoose Herpestes ichneumon http://maps.iucnredlist.org/map.html?id=41613 Portugal 5/45 (11.1) PCR Cumbassá et al., 2015 Herpestidae Indian grey mongoose Herpestes edwardsii http://maps.iucnredlist.org/map.html?id=41611 India n.a. (0.0) CAT Yadav and Sethi, 1980 n.a. CFT Enright et al., 1971 Spotted skunk Spilogale gracilis http://maps.iucnredlist.org/map.html?id=136797 USA Mephitidae n.a. SERO McQuiston et al., 2002 Striped skunk Mephitis mephitis http://maps.iucnredlist.org/map.html?id=41635 USA n.a. CFT Enright et al., 1971 American mink Neovison vison http://maps.iucnredlist.org/map.html?id=41661 Spain 0/3 (0.0) PCR Astobiza et al., 2011 Portugal 0/1 (0.0) PCR Cumbassá et al., 2015 Eurasian badger Meles meles http://maps.iucnredlist.org/map.html?id=29673 Spain 0/74 (0.0) PCR Astobiza et al., 2011 Carnivora Pine marten Martes martes http://maps.iucnredlist.org/map.html?id=12848 Spain 0/12 (0.0) PCR Astobiza et al., 2011 0/103 (0.0) PCR Duncan et al., 2015 Mustelidae Sea otter Enhydra lutris http://maps.iucnredlist.org/map.html?id=7750 USA 21/105 (20.0) IFA Duncan et al., 2015 0/33 (0.0) CFT White et al., 2013 Stone marten Martes foina http://maps.iucnredlist.org/map.html?id=29672 Spain 0/25 (0.0) PCR Astobiza et al., 2011 Portugal 0/2 (0.0) PCR Cumbassá et al., 2015 Least weasel Mustela nivalis http://maps.iucnredlist.org/map.html?id=14021 Spain 0/5 (0.0) PCR Astobiza et al., 2011 0/40 (0.0) PCR Minor et al., 2013 0/400 (0.0) PCR Duncan et al., 2014 Northern fur seal Callorhinus ursinus http://maps.iucnredlist.org/map.html?id=3590 USA Otariidae 109/146 (74.6) PCR Duncan et al., 2012 148/236 (62.7) IFA Minor et al., 2013 Steller sea lion Eumetopias jubatus http://maps.iucnredlist.org/map.html?id=8239 USA 44/74 (59.4) IFA Minor et al., 2013

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference Otariidae Steller sea lion Eumetopias jubatus http://maps.iucnredlist.org/map.html?id=8239 USA 1/2 (50.0) PCR Kersh et al., 2012

USA / 17/27 (63.0) PCR Kersh et al., 2012 Phocidae Harbor seal Phoca vitulina richardsi http://maps.iucnredlist.org/map.html?id=17013 Canada 73/215 (34.0) IFA Kersh et al., 2012 USA 2/26 (7.7) CFT Enright et al., 1971 Canada n.a. (7.1) IFA Marrie et al., 1993 Japan 0/559 (0.0) SERO Inoue et al., 2011 Procyonidae Northern raccon Procyon lotor http://maps.iucnredlist.org/map.html?id=41686 Spain 15/48 (31.5) PCR González-Barrio et al., unpublished 5/11 (45.4) SERO Randhawa et al. 1977 Carnivora USA n.a. SERO McQuiston et al., 2002 Brown bear Ursus arctos http://maps.iucnredlist.org/map.html?id=41688 Croatia 0/13 (0.0) CFT Madic et al., 1993 3/37 (8.1) CFT Dunbar et al., 1998 Ursidae American black bear Ursus americanus http://maps.iucnredlist.org/map.html?id=41687 USA 25/149 (17) MAT Ruppanner et al., 1982 Japanese black bear Ursus thibetanus http://maps.iucnredlist.org/map.html?id=22824 Japan 28/36 (77.7) ELISA Ejercito et al., 1993 Portugal 0/3 (0.0) PCR Cumbassá et al., 2015 Common genet Genetta genetta http://maps.iucnredlist.org/map.html?id=41698 Viverridae Spain 0/12 (0.0) PCR Astobiza et al., 2011 Masked palm civet Paguma larvata http://maps.iucnredlist.org/map.html?id=41692 Japan 0/10 (0.0) ELISA Ejercito et al., 1993 Cetacea Phocoenidae Harbour porpoise Phocoena phocoena http://maps.iucnredlist.org/map.html?id=17027 USA 2/6 (33.3) PCR Kersh et al., 2012 Molossidae Pallas's mastiff bat Molossus molossus http://maps.iucnredlist.org/map.html?id=13648 Fr. Guiana 0/57 (0.0) ELISA Gardon et al., 2001 Greater spear-nosed Chiroptera Phyllostomidae Phyllostomus hastatus http://maps.iucnredlist.org/map.html?id=17218 Fr. Guiana 0/17 (0.0) ELISA Gardon et al., 2001 bat Pteropodidae Indian flying fox Pteropus giganteus http://maps.iucnredlist.org/map.html?id=18725 India 2/14 (14.3) CAT Yadav and Sethi, 1980 Common opossum Didelphis marsupialis http://maps.iucnredlist.org/map.html?id=40501 Fr. Guiana 1/4 (25.0) ELISA Gardon et al., 2001 Didelphimorphia Didelphidae Gray four-eyed Philander opossum http://maps.iucnredlist.org/map.html?id=40516 Fr. Guiana 4/36 (11.1) ELISA Gardon et al., 2001 opossum Agile wallaby Macropus agilis http://maps.iucnredlist.org/map.html?id=40560 Australia 1/5 (20.0) ELISA Cooper et al., 2013

Diprotodontia Macropodidae Common walloroo Macropus robustus http://maps.iucnredlist.org/map.html?id=40565 Australia 1/3 (33.3) ELISA Cooper et al., 2013 Eastern grey Macropus giganteus http://maps.iucnredlist.org/map.html?id=41513 Australia n.a. (12.0) CFT Pope et al., 1960 kangaroo

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference n.a. (33.0) CFT Pope et al., 1960 Red kangaroo Macropus rufus http://maps.iucnredlist.org/map.html?id=40567 Australia 1/4 (25.0) ELISA Cooper et al., 2013 Black-striped wallaby Macropus dorsalis http://maps.iucnredlist.org/map.html?id=40562 Australia 1/1 (100.0) ELISA Cooper et al., 2013 Macropodidae 115/343 (33.5) ELISA Banazis et al., 2010

Diprotodontia Westerm grey 245/1017 (24.1) ELISA Potter et al., 2011 Macropus fuliginosus http://maps.iucnredlist.org/map.html?id=40563 Australia kangaroo 42/1017 (4.1) PCR Potter et al., 2011 42/343 (12.2) PCR Banazis et al., 2010 Phalangeridae Brushtail possum Trichosurus vulpecula http://maps.iucnredlist.org/map.html?id=40585 Australia 1/2 (50.0) ELISA Cooper et al., 2013 Potoroidae Rufous bettong Aepyprymnus rufescens http://maps.iucnredlist.org/map.html?id=40558 Australia 0/1 (0.0) ELISA Cooper et al., 2013 Southern white- Erinaceomorpha Erinaceidae Erinaceus concolor http://maps.iucnredlist.org/map.html?id=40605 Iran n.a. (0.0) SERO/PCR Mostafavi et al., 2012 breasted hedgehog Crowned shrew Sorex coronatus http://maps.iucnredlist.org/map.html?id=29663 Spain 0/14 (0.0) PCR Barandika et al., 2008 Soricidae Eulipotyphla White-toothed shrew Crocidura russula http://maps.iucnredlist.org/map.html?id=29652 Spain 0/16 (0.0) PCR Barandika et al., 2008 Talpidae European mole Talpa europaea http://maps.iucnredlist.org/map.html?id=41481 Spain 0/24 (0.0) PCR Barandika et al., 2008 Brush rabbit Sylvilagus bachmani http://maps.iucnredlist.org/map.html?id=41302 USA n.a. (53.0) CFT Enright et al., 1971 Czech Rep. 12/263 (4.5) MAT Rehacek et al., 1977 Czech Rep. 0/23 (0.0) MAT Hubalez et al., 1993 European hare Lepus europaeus http://maps.iucnredlist.org/map.html?id=41280 Cyprus 15/31 (48.4) PCR Psaroulaki et al., 2014 Spain 2/22 (9.1) PCR Astobiza et al., 2011 0/6 (0.0) PCR Astobiza et al., 2011 Lagomorpha Leporidae European rabbit Oryctolagus cuniculus http://maps.iucnredlist.org/map.html?id=41291 Spain 176/464 (37.9) ELISA González-Barrio et al., 2015b 6/136 (4.4) PCR González-Barrio et al., 2015b White-tailed Lepus townsendii http://maps.iucnredlist.org/map.html?id=41288 USA n.a. CFT Enright et al., 1971 jackrabbit Japanese hare Lepus brachyurus http://maps.iucnredlist.org/map.html?id=41275 Japan 5/8 (62.5) ELISA Ejercito et al., 1993 n.a. (49.0) IFA Marrie et al., 1993 Snowshoe hare Lepus americanus http://maps.iucnredlist.org/map.html?id=41273 Canada 11/22 (50.0) ELISA Marrie et al., 1986 Northern brown Peramelemorphia Peramelidae Isoodon macrourus http://maps.iucnredlist.org/map.html?id=40552 Australia 6/35 (17.1) ELISA Cooper et al., 2013 bandicoot

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference Pale-throated three- French Bradypodidae Bradypus tridactylus http://maps.iucnredlist.org/map.html?id=3037 1/1 (100.0) PCR Davoust et al., 2014 Pilosa toed sloth Guiana n.a. Ant eater n.a. India n.a. (0.0) CAT Yadav and Sethi, 1980 Primates Cercopithecidae Japanese macaque Macaca fuscata http://maps.iucnredlist.org/map.html?id=12552 Japan 15/54 (27.7) ELISA Ejercito et al., 1993 1/95 (1.0) MAT Rehacek et al., 1977

Czech Rep. 2/65 (3.0) CFT Syrucek and Raska, 1956

n.a. No ISO Syrucek and Raska, 1956

Bank vole Myodes glareolus http://maps.iucnredlist.org/map.html?id=4973 Italy 0/42 (0.0) PCR Pascucci et al., 2015

Slovakia 0/23 (0.0) PCR Smetanova et al., 2006

Spain 0/6 (0.0) PCR Barandika et al., 2008

UK 31/180 (17.2) ELISA Meredith et al., 2014

1/2 (50.0) CFT Enright et al., 1969

24/78 (30.0) CFT Enright et al., 1971 Brush mouse Peromyscus boylii http://maps.iucnredlist.org/map.html?id=16652 USA 5/306 (1.6) MAT Rieman et al., 1979

Rodentia Cricetidae n.a. No ISO Enright et al., 1971

Canyon mouse Peromyscus crinitus http://maps.iucnredlist.org/map.html?id=16656 USA 0/40 (0.0) CFT Stoenner et al., 1960

0/31 (0.0) MAT Rehacek et al., 1977

Czech Rep. 1/82 (1.2) CFT Syrucek and Raska, 1956

Common vole Microtus arvalis http://maps.iucnredlist.org/map.html?id=13488 n.a. No ISO Syrucek and Raska, 1956

Germany 0/119 (0.0) PCR Pluta et al., 2010

Slovakia 0/3 (0.0) PCR Smetanova et al., 2006

Canada n.a. (76.1) PCR Thompson et al., 2012

North American Peromyscus 0/364 (0.0) CFT Stoenner et al., 1960 http://maps.iucnredlist.org/map.html?id=16672 deermouse maniculatus USA 5/306 (1.6) MAT Rieman et al., 1979

6/24 (25.0) CFT Enright et al., 1969

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference CFT Enright et al., 1971 North American Peromyscus http://maps.iucnredlist.org/map.html?id=16672 USA 72/291 (24.7) No ISO Enright et al., 1971 deermouse maniculatus ISO Stoenner et al., 1960

Desert wood rat Neotoma lepida http://maps.iucnredlist.org/map.html?id=14589 USA 0/107 (0.0) CFT Stoenner et al., 1960

1/21 (4.7) MAT Rieman et al., 1979

2/31 (6.4) CFT Enright et al., 1969 Dusky-footed wood Neotoma fuscipes http://maps.iucnredlist.org/map.html?id=14587 USA 9/311 (2.9) CFT Enright et al., 1971 rat n.a. ISO Burgdorfer et al., 1963

n.a. No ISO Enright et al., 1971

European pine vole Microtus subterraneus http://maps.iucnredlist.org/map.html?id=13489 Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956

1/6 (16.6) CFT Syrucek and Raska, 1956 European water vole Arvicola amphibius http://maps.iucnredlist.org/map.html?id=2149 Czech Rep. n.a. No ISO Syrucek and Raska, 1956 Rodentia Cricetidae 0/11 (0.0) MAT Rehacek et al., 1977 Czech Rep. 0/3 (0.0) CFT Syrucek and Raska, 1956 Field vole Microtus agrestis http://maps.iucnredlist.org/map.html?id=13426 Poland n.a. ISO Tylewska-Wierzbanowska et al., 1991

UK 59/309 (19.1) ELISA Meredith et al., 2014

Czech Rep. n.a. ISO Proreshnaya et al., 1960

Gray dwarf hamster Cricetulus migratorius http://maps.iucnredlist.org/map.html?id=5528 Former 0/1 (0.0) CFT Yevdoshenko and Proreshnaya, 1961 USSR n.a. ISO Yevdoshenko and Proreshnaya, 1961

0/8 (0.0) MAT Rieman et al., 1979

Southern marsh Reithrodontomys 3/28 (10.7) CFT Enright et al., 1971 http://maps.iucnredlist.org/map.html?id=19410 USA harvest mouse megalotis 3/28 (10.7) CFT Stoenner et al., 1960

n.a. No ISO Enright et al., 1971 Microtus Chihuahua vole http://maps.iucnredlist.org/map.html?id=13452 USA 0/12 (0.0) MAT Rieman et al., 1979 pennsylvanicus

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference 1/5 (20.0) CFT Enright et al., 1969 Microtus Chihuahua vole http://maps.iucnredlist.org/map.html?id=13452 USA 20/100 (20.0) CFT Enright et al., 1971 pennsylvanicus n.a. No ISO Enright et al., 1971

Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956

0/6 (0.0) CFT Yevdoshenko and Proreshnaya, 1961 Musk rat Ondatra zibethicus http://maps.iucnredlist.org/map.html?id=15324 Former USSR n.a. No ISO Yevdoshenko and Proreshnaya, 1961 Cricetidae USA 2/19 (10.5) MAT Rieman et al., 1979 Northern grasshopper Onychomys leucogaster http://maps.iucnredlist.org/map.html?id=15338 USA 1/5 (20.0) CFT Stoenner et al., 1960 mouse 12/128 (9.3) CFT Enright et al., 1971

Pinyon mouse Peromyscus truei http://maps.iucnredlist.org/map.html?id=16694 USA n.a. No ISO Enright et al., 1971

0/11 (0.0) CFT Stoenner et al., 1960 Rodentia Revillagigedo Island Myodes gapperi http://maps.iucnredlist.org/map.html?id=42617 Canada n.a. (18.0) PCR Thompson et al., 2012 ded-backed vole Small five-toed jerboa Allactaga elater http://maps.iucnredlist.org/map.html?id=853 Czech Rep. n.a. ISO Proreshnaya et al.,1961 Dipodidae Woodland jumping Napaeozapus insignis http://maps.iucnredlist.org/map.html?id=42612 Canada n.a. (83.3) PCR Thompson et al., 2012 mouse Cuvier's/Cayenne Proechimys cuvieri/ Echimyidae http://maps.iucnredlist.org/map.html?id=22712110 Fr. Guiana 4/26 (15.4) IFA Gardon et al., 2001 spiny rat cayenne

North American 0/3 (0.0) CFT Stoenner et al., 1960 Erethizontidae Erethizon dorsatum http://maps.iucnredlist.org/map.html?id=8004 USA porcupine 0/6 (0.0) MAT Rieman et al., 1979 Muscardinus Hazel dormouse http://maps.iucnredlist.org/map.html?id=13992 Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956 avellanarius Gliridae Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956 Garden dormouse Eliomys quercinus http://maps.iucnredlist.org/map.html?id=7618 Spain n.a. ISO Perez Gallardo et al., 1952

Houserock chisel- 5/189 (2.6) CFT Stoenner et al., 1960 Heteromyidae Dipodomys microps http://maps.iucnredlist.org/map.html?id=42603 USA toothed kangaroo rat n.a. ISO Stoenner et al., 1960

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference Owyhee river Microdipodops http://maps.iucnredlist.org/map.html?id=42606 USA 0/2 (0.0) CFT Stoenner et al., 1960 kangaroo mouse megacephalus

Great-basin pocket 0/14 (0.0) CFT Stoenner et al., 1960 Perognathus parvus http://maps.iucnredlist.org/map.html?id=42610 USA mouse 0/22 (0.0) MAT Rieman et al., 1979

0/17 (0.0) MAT Rieman et al., 1979

Heermani's kangaroo 0/2 (0.0) CFT Enright et al., 1969 Heteromyidae Dipodomys heermanni http://maps.iucnredlist.org/map.html?id=42600 USA rat 1/59 (1.7) CFT Enright et al., 1971

n.a. No ISO Enright et al., 1971 Long-tailed pocket Chaetodipus formosus http://maps.iucnredlist.org/map.html?id=4331 USA 0/47 (0.0) CFT Stoenner et al., 1960 mouse 16/312 (5.1) CFT Stoenner et al., 1960 Ord's kangaroo rat Dipodomys ordii http://maps.iucnredlist.org/map.html?id=6691 USA n.a. ISO Stoenner et al., 1960

Lesser bandicoot rat Bandicota bengalensis http://maps.iucnredlist.org/map.html?id=2540 India n.a. ISO Yadav and Sethi, 1980 Rodentia 1/15 (6.6) CFT Syrucek and Raska, 1956 Czech Rep. n.a. No ISO Syrucek and Raska, 1956

Black rat Rattus rattus http://maps.iucnredlist.org/map.html?id=19360 Fr. Guiana 0/17 (0.0) ELISA Gardon et al., 2001

0/56 (0.0) ELISA Reusken et al., 2011 Netherl. 5/166 (3.0) PCR Reusken et al., 2011

Muridae 32/136 (23.5) PCR Psaroulaki et al., 2014 Brown / black rat R. norvegicus/R. rattus Cyprus 63/494 (12.7) IFA Psaroulaki et al., 2010

41/286 (14.3) CFT Syrucek and Raska, 1956 Czech Rep. n.a. ISO Syrucek and Raska, 1956

Brown rat Rattus norvegicus http://maps.iucnredlist.org/map.html?id=19353 Germany 7/524 (1.3) PCR Runge et al., 2013

2/21 (9.5) ISO Yadav and Sethi, 1980 India 3/21 (14.3) CAT Yadav and Sethi, 1980

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference Japan 0/54 (0.0) ELISA Ejercito et al., 1993

23/146 (15.7) ELISA Reusken et al., 2011 Netherl. Brown rat Rattus norvegicus http://maps.iucnredlist.org/map.html?id=19353 8/164 (4.8) PCR Reusken et al., 2011 UK n.a. (7.0-53.0) ELISA Webster et al., 1995

USA 3/30 (10%) MAT Rieman et al., 1979

Great gerbil Rhombomys opimus http://maps.iucnredlist.org/map.html?id=19686 F.USSR n.a. ISO Zhmaeva et al., 1955

1/78 (1.3) MAT Rehacek et al., 1977

14/155 (9) CFT Syrucek and Raska, 1956 Czech Rep. n.a. ISO Proreshnaya et al., 1960

n.a. ISO Syrucek and Raska, 1956

1/6 (16.6) CFT Yevdoshenko and Proreshnaya, 1961 F. USSR House mouse Mus musculus http://maps.iucnredlist.org/map.html?id=13972 n.a. ISO Yevdoshenko and Proreshnaya, 1961 Rodentia Muridae Fr. Guiana 0/58 (0.0) IFA Gardon et al., 2001

0/4 (0.0) CAT Yadav and Sethi, 1980 India n.a. No ISO Yadav and Sethi, 1980

Spain 2/28 (7.1) PCR Barandika et al., 2008

USA 0/83 (0.0) MAT Rieman et al., 1979

Rat Rattus spp. http://maps.iucnredlist.org/map.html?id=19360 Spain 3/3 (100.0) PCR Jado et al., 2012

Shaw's jird Meriones shawi http://maps.iucnredlist.org/map.html?id=42666 Morocco n.a. ISO Blanc et al., 1947

0/16 (0.0) CFT Yevdoshenko and Proreshnaya, 1961 Tamarisk gerbil Meriones tamariscinus http://maps.iucnredlist.org/map.html?id=13169 F. USSR n.a. No ISO Yevdoshenko and Proreshnaya, 1961

Czech Rep. n.a. No ISO Syrucek and Raska, 1956 Long-tailed field Apodemus sylvaticus http://maps.iucnredlist.org/map.html?id=1904 Italy 2/101 (19.8) PCR Pascucci et al., 2015 mouse Slovakia 0/3 (0.0) PCR Smetanova et al., 2006

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference

Long-tailed field Spain 1/162 (0.6) PCR Barandika et al., 2008 Apodemus sylvaticus http://maps.iucnredlist.org/map.html?id=1904 mouse UK 48/307 (15.6) ELISA Meredith et al., 2014 1/202 (0.5) MAT Rehacek et al., 1977 Czech Rep. 11/212 (5.2) CFT Syrucek and Raska, 1956 Yellow-necked field Muridae Apodemus flavicollis http://maps.iucnredlist.org/map.html?id=1892 mouse Slovakia 1/38 (2.6) PCR Smetanova et al., 2006

Spain 0/3 (0.0) PCR Barandika et al., 2008

Barbary striped grass Kenya n.a. ISO Heisch, 1960 Lemniscomys barbarus http://maps.iucnredlist.org/map.html?id=11487 mouse Morocco n.a. ISO Blanc and Bruneau, 1956 Myocastoridae Coypu Myocastor coypus http://maps.iucnredlist.org/map.html?id=14085 Japan 4/32 (12.5) ELISA Ejercito et al., 1993 n.a. ISO Burgdorfer et al., 1963 Yellow-pine chipmunk Tamias amoenus http://maps.iucnredlist.org/map.html?id=42569 USA 1/85 (1.2) MAT Rieman et al., 1979 19/145 (13.1) CFT Enright et al., 1971

California ground 6/26 (23.0) CFT Enright et al., 1969 Rodentia Spermophilus beecheyi http://maps.iucnredlist.org/map.html?id=20481 USA squirrel 7/112 (6.2) MAT Rieman et al., 1979 n.a. ISO Enright et al., 1971 Arizona black-tailed Cynomys ludovicianus http://maps.iucnredlist.org/map.html?id=6091 USA n.a. CFT Enright et al., 1971 prairie dog Cliff chipmunk Tamias dorsalis http://maps.iucnredlist.org/map.html?id=42571 USA 0/2 (0.0) CFT Stoenner et al., 1960 Sciuridae Colorado chipmunk Tamias quadrivittatus http://maps.iucnredlist.org/map.html?id=42576 USA 0/1 (0.0) CFT Stoenner et al., 1960 Douglas squirrel Tamiasciurus douglasii http://maps.iucnredlist.org/map.html?id=42586 USA 2/3 (66.6) MAT Rieman et al., 1979 Eastern chipmunks Tamias striatus http://maps.iucnredlist.org/map.html?id=42583 Canada 0/12 (0.0) PCR Thompson et al., 2012

Golden-mantled 0/34 (0.0) MAT Rieman et al., 1979 Spermophilus lateralis http://maps.iucnredlist.org/map.html?id=42468 USA ground squirrel n.a. ISO Burgdorfer et al., 1963 4/72 (5.5) CFT Enright et al., 1971 Eastern gray squirrel Sciurus carolinensis http://maps.iucnredlist.org/map.html?id=42462 USA n.a. No ISO Enright et al., 1971 New Mexico least Tamias minimus http://maps.iucnredlist.org/map.html?id=42572 1/8 (12.5) CFT Stoenner et al., 1960 chipmunk

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference Long-clawed ground Spermophilopsis Former http://maps.iucnredlist.org/map.html?id=20471 n.a. ISO Zhmaeva et al., 1955 squirrel leptodactylus USSR Tamiasciurus Red squirrel http://maps.iucnredlist.org/map.html?id=42587 Canada n.a. (40.0) PCR Thompson et al., 2012 hudsonicus Carolina flying Glaucomys sabrinus http://maps.iucnredlist.org/map.html?id=39553 Canada n.a. (38.0) PCR Thompson et al., 2012 squirrel Eurasian red squirrel Sciurus vulgaris http://maps.iucnredlist.org/map.html?id=20025 Czech Rep. 0/2 (0.0) CFT Syrucek and Raska, 1956 Czech Rep. n.a. ISO Proreshnaya et al., 1960 Tien Shan ground Former Spermophilus relictus http://maps.iucnredlist.org/map.html?id=20491 12/182 (6.6) CFT Yevdoshenko and Proreshnaya, 1961 squirrel USSR Czech Rep. n.a. ISO Yevdoshenko and Proreshnaya, 1961 Rodentia Sciuridae Rock squirrel Spermophilus variegatus http://maps.iucnredlist.org/map.html?id=20495 USA 0/1 (0%) CFT Stoenner et al., 1960 2/9 (22.2%) CFT Enright et al., 1971 Sonoma chipmunk Tamias sonomae http://maps.iucnredlist.org/map.html?id=42581 USA n.a. No ISO Enright et al., 1971 Southern flying Glaucomys volans http://maps.iucnredlist.org/map.html?id=9240 Canada n.a. (65.0) PCR Thompson et al., 2012 squirrel Townsend's ground Spermophilus http://maps.iucnredlist.org/map.html?id=20476 USA 0/1 (0.0) CFT Stoenner et al., 1960 squirrel townsendii Western gray squirrel Sciurus griseus http://maps.iucnredlist.org/map.html?id=20011 USA 0/1 (0.0) MAT Rieman et al., 1979 White-tailed antelope Ammospermophilus http://maps.iucnredlist.org/map.html?id=42452 USA 0/104 (0.0) CFT Stoenner et al., 1960 squirrel leucurus Asian house shrew Suncus murinus http://maps.iucnredlist.org/map.html?id=41440 India 1/24 (4.1) CAT Yadav and Sethi, 1980 Bicoloured white- Crocidura leucodon http://maps.iucnredlist.org/map.html?id=29651 Czech Rep. 0/2 (0.0) MAT Rehacek et al., 1977 Soricomorpha Soricidae toothed shrew Common shrew Sorex araneus http://maps.iucnredlist.org/map.html?id=29661 Czech Rep. 3/14 (21.4) MAT Rehacek et al., 1977 Eurasian pygmy shrew Sorex minutus http://maps.iucnredlist.org/map.html?id=29667 Czech Rep. 0/3 (0.0) MAT Rehacek et al., 1977

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference Black kite Milvus migrans http://maps.iucnredlist.org/map.html?id=22734972 Spain 1/7 (14.0) PCR Astobiza et al., 2011 1/1 (100.0) PCR Psaroulaki et al., 2014* Bonelli's eagle Aquila fasciata http://maps.iucnredlist.org/map.html?id=22696076 Cyprus 1/2 (50.0) PCR Ioannou et al., 2009* Brahminy kite Haliastur indus http://maps.iucnredlist.org/map.html?id=22695094 India n.a. (0.0) CAT Yadav and Sethi, 1980 1/2 (50.0) PCR Psaroulaki et al., 2014 Eurasian buzzard Buteo buteo http://maps.iucnredlist.org/map.html?id=61695117 Cyprus 1/4 (25.0) PCR Ioannou et al., 2009 Eurasian griffon Portugal 0/19 (0.0) PCR Cumbassá et al., 2015 Gyps fulvus http://maps.iucnredlist.org/map.html?id=22695219 vulture Spain 1/9 (11.0) PCR Astobiza et al., 2011 Eurasian 0/1 (0.0) PCR Psaroulaki et al., 2014 Accipiter nisus http://maps.iucnredlist.org/map.html?id=22695624 Cyprus sparrowhawk 0/2 (50.0) PCR Ioannou et al., 2009 Accipitriformes Accipitridae European honey 1/3 (33.3) PCR Ioannou et al., 2009 Pernis apivorus http://maps.iucnredlist.org/map.html?id=22694989 Cyprus buzzard 4/4 (100.0) PCR Psaroulaki et al., 2014 1/1 (100.0) PCR Psaroulaki et al., 2014 Hen harrier Circus cyaneus http://maps.iucnredlist.org/map.html?id=22727733 Cyprus 2/3 (66.6) PCR Ioannou et al., 2009 1/2 (50.0) PCR Psaroulaki et al., 2014 Cyprus Long-legged buzzard Buteo rufinus http://maps.iucnredlist.org/map.html?id=22736562 1/3 (66.6) PCR Ioannou et al., 2009 India n.a. (0.0) CAT Yadav and Sethi, 1980 1/1 (100.0) PCR Psaroulaki et al., 2014 Northern goshawk Accipiter gentilis http://maps.iucnredlist.org/map.html?id=22695683 Cyprus 3/3 (100.0) PCR Ioannou et al., 2009 Red-tailed hawk Buteo jamaicensis http://maps.iucnredlist.org/map.html?id=22695933 USA 1/1 (100.0) MAT Rieman et al., 1979 Western marsh harrier Circus aeruginosus http://maps.iucnredlist.org/map.html?id=22695344 Cyprus 0/5 (0.0) PCR Psaroulaki et al., 2014 Canada goose Branta canadensis http://maps.iucnredlist.org/map.html?id=22679935 USA 0/1 (0.0) MAT Rieman et al., 1979 Cinnamon teal Spatula cyanoptera http://maps.iucnredlist.org/map.html?id=22680233 USA 1/4 (25.0) MAT Rieman et al., 1979 0/1 (0.0) PCR Ioannou et al., 2009 Common teal Anas crecca http://maps.iucnredlist.org/map.html?id=22729717 Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014 Gadwall Mareca strepera http://maps.iucnredlist.org/map.html?id=22680149 USA 1/1 (100.0) MAT Rieman et al., 1979 1/1 (100.0) PCR Ioannou et al., 2009 Cyprus 1/2 (50.0) PCR Psaroulaki et al., 2014 India n.a. (0.0) CAT Yadav and Sethi, 1980 Anseriformes Anatidae Mallard Anas platyrhynchos http://maps.iucnredlist.org/map.html?id=22680186 n.a. (0.0) PCR To et al., 1998 Japan 0/101 (0.0) MAT To et al., 1998 Spain 0/3 (0.0) PCR Astobiza et al., 2011 USA 1/14 (7.1) MAT Rieman et al., 1979 Pintail Anas acuta http://maps.iucnredlist.org/map.html?id=22680301 USA 0/5 (0.0) MAT Rieman et al., 1979 Tundra swan Cygnus columbianus http://maps.iucnredlist.org/map.html?id=22679862 USA 1/1 (100.0) MAT Rieman et al., 1979 n.a. (0.0) PCR To et al., 1998 Whooper swan Cygnus cygnus http://maps.iucnredlist.org/map.html?id=22679856 Japan 0/10 (0.0) MAT To et al., 1998 0/1 (0.0) PCR Ioannou et al., 2009 Bucerotiformes Upupidae Common hoopoe Upupa epops http://maps.iucnredlist.org/map.html?id=22682655 Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference Caprimulgiformes Caprimulgidae European nightjar Caprimulgus europaeus http://maps.iucnredlist.org/map.html?id=22689887 Cyprus 1/1 (100.0) PCR Psaroulaki et al., 2014 Cathartiformes Cathartidae Turkey vulture Cathartes aura http://maps.iucnredlist.org/map.html?id=22697627 USA 2/4 (50.0) MAT Rieman et al., 1979 Alcidae n.a. n.a. Spain 0/3 (0.0) PCR Astobiza et al., 2011 Burhinidae n.a. n.a. Spain 0/1 (0.0) PCR Astobiza et al., 2011 Charadriidae n.a. n.a. Spain 0/5 (0.0) PCR Astobiza et al., 2011 Laridae n.a. n.a. Spain 0/1 (0.0) PCR Astobiza et al., 2011 0/1 (0.0) PCR Ioannou et al., 2009 Burhinidae Eurasian thick-knee Burhinus oedicnemus http://maps.iucnredlist.org/map.html?id=45111439 Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014 0/1 (0.0) PCR Ioannou et al., 2009 Black-headed gull Larus ridibundus http://maps.iucnredlist.org/map.html?id=22694420 Cyprus Charadriiformes 0/2 (0.0) PCR Psaroulaki et al., 2014 1/4 (25.0) PCR Ioannou et al., 2009 Laridae Caspian gull Larus cachinnans http://maps.iucnredlist.org/map.html?id=22735929 Cyprus 2/3 (66.6) PCR Psaroulaki et al., 2014 Lesser black-backed 1/1 (100.0) PCR Psaroulaki et al., 2014 Larus fuscus http://maps.iucnredlist.org/map.html?id=22694373 Cyprus gull 1/2 (50.0) PCR Ioannou et al., 2009 0/1 (0.0) PCR Ioannou et al., 2009 Recurvirostridae Black-winged stilt Himantopus himantopus http://maps.iucnredlist.org/map.html?id=22727969 Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014 Scolopacidae Eurasian woodcock Scolopax rusticola http://maps.iucnredlist.org/map.html?id=22693052 Cyprus 0/2 (0.0) PCR Ioannou et al., 2009 1/1 (100.0) PCR Ioannou et al., 2009 Cyprus Ciconiiformes Ciconiidae White stork Ciconia ciconia http://maps.iucnredlist.org/map.html?id=22697691 1/1 (100.0) PCR Psaroulaki et al., 2014 Spain 0/5 (0.0) PCR Astobiza et al., 2011 Band-tailed pigeon Patagioenas fasciata http://maps.iucnredlist.org/map.html?id=22725264 USA 3/31 (9.6) MAT Rieman et al., 1979 3/10 (30.0) PCR Ioannou et al., 2009 Common woodpigeon Columba palumbus http://maps.iucnredlist.org/map.html?id=22690103 Cyprus 8/15 (53.3) PCR Psaroulaki et al., 2014 European turtle-dove Streptopelia turtur http://maps.iucnredlist.org/map.html?id=22690419 Cyprus 1/7 (14.3) PCR Ioannou et al., 2009 Laughing turtle-Dove Spilopelia senegalensis http://maps.iucnredlist.org/map.html?id=22690445 India n.a. (0.0) CAT Yadav and Sethi, 1980 Columbiformes Columbidae Mourning dove Zenaida macroura http://maps.iucnredlist.org/map.html?id=22690736 USA 0/9 (0.0) MAT Rieman et al., 1979 India 1/11 (9.0) CAT Yadav and Sethi, 1980 Iran n.a. (7.9) n.a. Mostafavi et al., 2012 Rock dove Columba livia http://maps.iucnredlist.org/map.html?id=22690066 France n.a. ISO Stein et al., 1999 n.a. (0.0) PCR To et al., 1998 Japan 4/100 (4.0) MAT To et al., 1998 0/1 (0.0) PCR Psaroulaki et al., 2014 Alcedinidae Common kingfisher Alcedo atthis http://maps.iucnredlist.org/map.html?id=22683027 Cyprus 0/2 (0.0) PCR Ioannou et al., 2009 Coraciiformes 0/2 (0.0) PCR Ioannou et al., 2009 Meropidae European bee-eater Merops apiaster http://maps.iucnredlist.org/map.html?id=22683756 Cyprus 0/2 (0.0) PCR Psaroulaki et al., 2014

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference Acciprinidae Indian vulture Gyps indicus http://maps.iucnredlist.org/map.html?id=22729731 India n.a. (0.0) CAT Yadav and Sethi, 1980 n.a. n.a. Spain 0/15 (0.0) PCR Astobiza et al., 2011 4/11 (36.3) PCR Ioannou et al., 2009 Cyprus Common kestrel Falco tinnunculus http://maps.iucnredlist.org/map.html?id=22696362 9/19 (47.3) PCR Psaroulaki et al., 2014 Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956 0/1 (0.0) PCR Ioannou et al., 2009 Eleonora's falcon Falco eleonorae http://maps.iucnredlist.org/map.html?id=22696442 Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014 Falcnoniformes 0/1 (0.0) PCR Ioannou et al., 2009 Falconidae Eurasian hobby Falco subbuteo http://maps.iucnredlist.org/map.html?id=22696460 Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014 1/1 (100.0) PCR Psaroulaki et al., 2014 Peregrine falcon Falco peregrinus http://maps.iucnredlist.org/map.html?id=45354964 Cyprus 2/4 (50.0) PCR Ioannou et al., 2009 0/1 (0.0) PCR Ioannou et al., 2009 Red-footed falcon Falco vespertinus http://maps.iucnredlist.org/map.html?id=22696432 Cyprus 0/2 (0.0) PCR Psaroulaki et al., 2014 0/1 (0.0) PCR Ioannou et al., 2009 Saker falcon Falco cherrug http://maps.iucnredlist.org/map.html?id=22696495 Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014 0/2 (0.0) PCR Ioannou et al., 2009 Black francolin Francolinus francolinus http://maps.iucnredlist.org/map.html?id=22678719 Cyprus 0/2 (0.0) PCR Psaroulaki et al., 2014 0/1 (0.0) PCR Psaroulaki et al., 2014 Chukar Alectoris chukar http://maps.iucnredlist.org/map.html?id=22678691 Cyprus 0/2 (0.0) PCR Ioannou et al., 2009 0/3 (0.0) PCR Ioannou et al., 2009 Common quail Coturnix coturnix http://maps.iucnredlist.org/map.html?id=22678944 Cyprus Phasianidae 1/2 (50.0) PCR Psaroulaki et al., 2014 Galliformes Grey partridge Perdix perdix http://maps.iucnredlist.org/map.html?id=22678911 Czech Rep. 0/5 (0.0) CFT Syrucek and Raska, 1956 Peacock n.a. India n.a. (0.0) CAT Yadav and Sethi, 1980 Guinea fowl n.a. India n.a. (0.0) CAT Yadav and Sethi, 1980 Czech Rep. 0/3 (0.0) CFT Syrucek and Raska, 1956 Common pheasant Phasianus colchicus http://maps.iucnredlist.org/map.html?id=45100023 Spain 0/5 (0.0) PCR Astobiza et al., 2011 Odontophoridae Mountain quail Oreortyx pictus http://maps.iucnredlist.org/map.html?id=22679591 USA 0/6 (0.0) MAT Rieman et al., 1979 1/1 (100.0) PCR Ioannou et al., 2009 Gruidae Common crane Grus grus http://maps.iucnredlist.org/map.html?id=22692146 Cyprus 1/1 (100.0) PCR Psaroulaki et al., 2014 American coot Fulica americana http://maps.iucnredlist.org/map.html?id=62169677 USA 15/33 (45.4) MAT Rieman et al., 1979 1/5 (20.0) PCR Ioannou et al., 2009 Common coot Fulica atra http://maps.iucnredlist.org/map.html?id=22692913 Cyprus 3/4 (75.0) PCR Psaroulaki et al., 2014 1/5 (20.0) PCR Ioannou et al., 2009 Gruiformes Common moorhen Gallinula chloropus http://maps.iucnredlist.org/map.html?id=62120190 Cyprus Rallidae 5/7 (71.4) PCR Psaroulaki et al., 2014 0/1 (0.0) PCR Psaroulaki et al., 2014 Corncrake Crex crex http://maps.iucnredlist.org/map.html?id=22692543 Cyprus 0/2 (0.0) PCR Ioannou et al., 2009 n.a. n.a. Spain 0/3 (0.0) PCR Astobiza et al., 2011 Western water rail Rallus aquaticus http://maps.iucnredlist.org/map.html?id=22725141 Cyprus 0/2 (0.0) PCR Ioannou et al., 2009

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference Aegithalidae American bushtit Psaltriparus minimus http://maps.iucnredlist.org/map.html?id=22712028 USA 0/3 (0.0) MAT Rieman et al., 1979 Alaudidae Eurasian skylark Alauda arvensis http://maps.iucnredlist.org/map.html?id=22717415 Czech Rep. 0/2 (0.0) CFT Syrucek and Raska, 1956 Certhiidae Eurasian treecreeper Certhia familiaris http://maps.iucnredlist.org/map.html?id=22735060 Czech Rep. 0/2 (0.0) CFT Syrucek and Raska, 1956 American crow Corvus brachyrhynchos http://maps.iucnredlist.org/map.html?id=22705990 USA 12/41 (29.2) MAT Rieman et al., 1979 Western scrub jay Aphelocoma californica http://maps.iucnredlist.org/map.html?id=22705623 USA 1/6 (16.6) MAT Rieman et al., 1979 Czech Rep. 0/2 (0.0) CFT Syrucek and Raska, 1956 India n.a. ISO Yadav and Sethi, 1980 Carrion crow Corvus corone http://maps.iucnredlist.org/map.html?id=22706016 13/173 (7.5.0) PCR To et al., 1998 Japan 64/173 (37.0) MAT To et al., 1998 Spain 0/6 (0.0) PCR Astobiza et al., 2011 Poland n.a. ISO Tylewska-Wierzbanowska et al., 1991 Corvidae Common raven Corvus corax http://maps.iucnredlist.org/map.html?id=22706068 USA 1/2 (50.0) MAT Rieman et al., 1979 Eurasian jay Garrulus glandarius http://maps.iucnredlist.org/map.html?id=22705764 Czech Rep. 0/5 (0.0) CFT Syrucek and Raska, 1956 5/41 (12.2) PCR To et al., 1998 Jungle crow Corvus macrorhynchos http://maps.iucnredlist.org/map.html?id=22706019 Japan 91/258 (35.3) MAT To et al., 1998 Black-billed magpie Pica pica hudsoni http://maps.iucnredlist.org/map.html?id=22705865 USA 0/3 (0.0) MAT Rieman et al., 1979 Rook Corvus frugilegus http://maps.iucnredlist.org/map.html?id=22706016 Czech Rep. 0/21 (0.0) CFT Syrucek and Raska, 1956 Passeriformes Spotted nutcracker Nucifraga caryocatactes http://maps.iucnredlist.org/map.html?id=22705912 Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956 Steller´s jay Cyanocitta stelleri http://maps.iucnredlist.org/map.html?id=22705614 USA 0/9 (0.0) MAT Rieman et al., 1979 Canyon towhee Melozone fuscus http://maps.iucnredlist.org/map.html?id=22721331 USA 0/4 (0.0) MAT Rieman et al., 1979 Golden-crowned Zonotrichia atricapilla http://maps.iucnredlist.org/map.html?id=22721091 USA 8/10 (80.0) MAT Rieman et al., 1979 sparrow Emberizidae Dark-eyed junco Junco hyemalis http://maps.iucnredlist.org/map.html?id=22735032 USA 1/2 (50.0) MAT Rieman et al., 1979 White-crowned Zonotrichia leucophrys http://maps.iucnredlist.org/map.html?id=22721088 USA 34/48 (70.8) MAT Rieman et al., 1979 sparrow Yellowhammer Emberiza citrinella http://maps.iucnredlist.org/map.html?id=22720878 Czech Rep. 1/29 (3.4) CFT Syrucek and Raska, 1956 Eurasian bullfinch Pyrrhula pyrrhula http://maps.iucnredlist.org/map.html?id=22720671 Czech Rep. 0/3 (0.0) CFT Syrucek and Raska, 1956 Eurasian chaffinch Fringilla coelebs http://maps.iucnredlist.org/map.html?id=22720030 Czech Rep. 1/45 (2.2) CFT Syrucek and Raska, 1956 Eurasian siskin Carduelis spinus http://maps.iucnredlist.org/map.html?id=22720354 Czech Rep. 0/8 (0.0) CFT Syrucek and Raska, 1956 European goldfinch Carduelis carduelis http://maps.iucnredlist.org/map.html?id=22720419 Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956 Fringillidae European greenfinch Carduelis chloris http://maps.iucnredlist.org/map.html?id=22720330 Czech Rep. 0/8 (0.0) CFT Syrucek and Raska, 1956 Island canary Serinus canaria http://maps.iucnredlist.org/map.html?id=22720056 Czech Rep. 0/3 (0.0) CFT Syrucek and Raska, 1956 House finch Carpodacus mexicanus http://maps.iucnredlist.org/map.html?id=22720563 USA 0/82 (0.0) MAT Rieman et al., 1979 Red crossbill Loxia curvirostra http://maps.iucnredlist.org/map.html?id=22720646 Czech Rep. 0/4 (0.0) CFT Syrucek and Raska, 1956

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference Barn swallow Hirundo rustica http://maps.iucnredlist.org/map.html?id=22712252 Czech Rep. 8/87 (9.2) CFT Syrucek and Raska, 1956 N. house martin Delichon urbicum http://www.iucnredlist.org/details/22712477/0 Czech Rep. 14/53 (26.4) CFT Syrucek and Raska, 1956 Hirundinidae India 1/200 (0.5) ISO Yadav and Sethi, 1980 Swallow n.a. India 6/200 (3.0) CAT Yadav and Sethi, 1980 Euphagus Brewer´s blackbird http://maps.iucnredlist.org/map.html?id=22724332 USA 6/18 (33.3) MAT Rieman et al., 1979 cyanocephalus Icteridae Western meadowlark Sturnella neglecta http://maps.iucnredlist.org/map.html?id=22724256 USA 1/2 (50.0) MAT Rieman et al., 1979 Red-winged blackbird Agelaius phoeniceus http://maps.iucnredlist.org/map.html?id=22724191 USA 0/4 (0.0) MAT Rieman et al., 1979 0/1 (0.0) PCR Ioannou et al., 2009 Lesser grey shrike Lanius minor http://maps.iucnredlist.org/map.html?id=22705038 Cyprus Laniidae 0/1 (0.0) PCR Psaroulaki et al., 2014 Red-backed shrike Lanius collurio http://maps.iucnredlist.org/map.html?id=22705001 Czech Rep. 0/2 (0.0) CFT Syrucek and Raska, 1956 Grey wagtail Motacilla cinerea http://maps.iucnredlist.org/map.html?id=22718392 Czech Rep. 0/2 (0.0) CFT Syrucek and Raska, 1956 Motacillidae Tree pipit Anthus trivialis http://maps.iucnredlist.org/map.html?id=22718546 Czech Rep. 0/2 (0.0) CFT Syrucek and Raska, 1956 White wagtail Motacilla alba http://maps.iucnredlist.org/map.html?id=22718348 Czech Rep. 1/15 (6.6) CFT Syrucek and Raska, 1956 Black redstart Phoenicurus ochruros http://maps.iucnredlist.org/map.html?id=22710051 Czech Rep. 2/22 (9.0) CFT Syrucek and Raska, 1956 Passeriformes Common redstart Phoenicurus phoenicurus http://maps.iucnredlist.org/map.html?id=22710055 Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956 Muscicapidae European robin Erithacus rubecula http://maps.iucnredlist.org/map.html?id=22709675 Czech Rep. 0/3 (0.0) CFT Syrucek and Raska, 1956 Spotted flycatcher Muscicapa striata http://maps.iucnredlist.org/map.html?id=22709192 Czech Rep. 0/7 (0.0) CFT Syrucek and Raska, 1956 Black-capped Parus atricapillus http://maps.iucnredlist.org/map.html?id=22711716 Czech Rep. 0/2 (0.0) CFT Syrucek and Raska, 1956 chickadee Blue tit Parus caeruleus http://maps.iucnredlist.org/map.html?id=22711944 Czech Rep. 0/3 (0.0) CFT Syrucek and Raska, 1956 Paridae Coal tit Parus ater http://maps.iucnredlist.org/map.html?id=22735965 Czech Rep. 0/7 (0.0) CFT Syrucek and Raska, 1956 Crested tit Parus cristatus http://maps.iucnredlist.org/map.html?id=22711810 Czech Rep. 0/6 (0.0) CFT Syrucek and Raska, 1956 Great tit Parus major http://maps.iucnredlist.org/map.html?id=22735990 Czech Rep. 1/7 (14.3) CFT Syrucek and Raska, 1956 Eurasian tree sparrow Passer montanus http://maps.iucnredlist.org/map.html?id=22718270 Czech Rep. 0/8 (0.0) CFT Syrucek and Raska, 1956 Czech Rep. 5/31 (16.1) CFT Syrucek and Raska, 1956 Passeridae House sparrow Passer domesticus http://maps.iucnredlist.org/map.html?id=22718174 India n.a. (0.0) CAT Yadav and Sethi, 1980 USA 7/14 (50.0) MAT Rieman et al., 1979 Ploceidae n.a. n.a. n.a. Spain 0/1 (0.0) PCR Astobiza et al., 2011 Reguliidae Goldcrest Regulus regulus http://maps.iucnredlist.org/map.html?id=22734997 Czech Rep. 0/2 (0.0) CFT Syrucek and Raska, 1956 19/69 (27.5) CAT Yadav and Sethi, 1980 Sturnidae Common myna Acridotheres tristis http://maps.iucnredlist.org/map.html?id=22710921 India 3/69 (4.3) ISO Yadav and Sethi, 1980

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference n.a. n.a. Spain 0/4 (0.0) PCR Astobiza et al., 2011 Sturnidae Czech Rep. 0/10 (0.0) CFT Syrucek and Raska, 1956 Common starling Sturnus vulgaris http://maps.iucnredlist.org/map.html?id=22710886 USA 18/157 (11.4) MAT Rieman et al., 1979 Common chiffchaff Phylloscopus collybita http://maps.iucnredlist.org/map.html?id=22715244 Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956 Icterine warbler Hippolais icterina http://maps.iucnredlist.org/map.html?id=2271491 Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956 Sylviidae n.a. n.a. Spain 0/1 (0.0) PCR Astobiza et al., 2011 Passeriformes Warbler Sylvia sp. Czech Rep. 0/10 (0.0) CFT Syrucek and Raska, 1956 Troglodytidae Winter wren Troglodytes troglodytes http://maps.iucnredlist.org/map.html?id=22711483 Czech Rep. 1/2 (50.0) CFT Syrucek and Raska, 1956 Eurasian blackbird Turdus merula http://maps.iucnredlist.org/map.html?id=2270877 Czech Rep. 0/2 (0.0) CFT Syrucek and Raska, 1956 Fieldfare Turdus pilaris http://maps.iucnredlist.org/map.html?id=22708816 Czech Rep. 0/2 (0.0) CFT Syrucek and Raska, 1956 Turdidae Mistle thrush Turdus viscivorus http://maps.iucnredlist.org/map.html?id=22708829 Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956 American robin Turdus migratorius http://maps.iucnredlist.org/map.html?id=22708958 USA 3/19 (15.8) MAT Rieman et al., 1979 Song thrush Turdus philomelos http://maps.iucnredlist.org/map.html?id=22708822 Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956 Black-crowned night- 1/1 (100.0) PCR Ioannou et al., 2009 Nycticorax nycticorax http://maps.iucnredlist.org/map.html?id=22697211 Cyprus heron 1/1 (100.0) PCR Psaroulaki et al., 2014 1/1 (100.0) PCR Ioannou et al., 2009 Common little bittern Ixobrychus minutus http://maps.iucnredlist.org/map.html?id=22735766 Cyprus 1/1 (100.0) PCR Psaroulaki et al., 2014 Eurasian bittern Botaurus stellaris http://maps.iucnredlist.org/map.html?id=22697346 Cyprus 0/1 (0.0) PCR Ioannou et al., 2009 1/2 (50.0) PCR Psaroulaki et al., 2014 Grey heron Ardea cinerea http://maps.iucnredlist.org/map.html?id=22696993 Cyprus Pelecaniformes Ardeidae 1/4 (25.0) PCR Ioannou et al., 2009 Indian pond heron Ardeola grayii http://maps.iucnredlist.org/map.html?id=22697128 India n.a. (0.0) CAT Yadav and Sethi, 1980 0/1 (0.0) PCR Ioannou et al., 2009 Little egret Egretta garzetta http://maps.iucnredlist.org/map.html?id=62774969 Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014 n.a. n.a. Spain 0/12 (0.0) PCR Astobiza et al., 2011 0/1 (0.0) PCR Ioannou et al., 2009 Purple heron Ardea purpurea http://maps.iucnredlist.org/map.html?id=22697031 Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014 1/2 (50.0) PCR Psaroulaki et al., 2014 Phoenicopteriformes Phoenicopteridae American flamingo Phoenicopterus ruber http://maps.iucnredlist.org/map.html?id=22729706 Cyprus 1/3 (33.3) PCR Ioannou et al., 2009 Great spotted Dendrocopos major http://maps.iucnredlist.org/map.html?id=22681124 Czech Rep. 0/7 (0.0) CFT Syrucek and Raska, 1956 woodpecker Piciformes Picidae Grey-faced Picus canus http://maps.iucnredlist.org/map.html?id=22726503 Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956 woodpecker 0/1 (0.0) PCR Ioannou et al., 2009 Great crested grebe Podiceps cristatus http://maps.iucnredlist.org/map.html?id=22696602 Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014 Podicipediformes Podicipedidae 0/1 (0.0) PCR Psaroulaki et al., 2014 Little grebe Tachybaptus ruficollis http://maps.iucnredlist.org/map.html?id=22696545 Cyprus 0/2 (0.0) PCR Ioannou et al., 2009 n.a. n.a. Spain 0/1 (0.0) PCR Astobiza et al., 2011 Procellariiformes Procellariidae n.a. n.a. Spain 0/3 (0.0) PCR Astobiza et al., 2011

Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference 1/56 (1.8) ISO Yadav and Sethi 1980 Psittaciformes Psittacidae Rose-ringed parakeet Psittacula krameri http://maps.iucnredlist.org/map.html?id=22685441 India 13/56 (23.2) CAT Yadav and Sethi 1980 Great horned owl Bubo virginianus http://maps.iucnredlist.org/map.html?id=61752071 USA 0/1 (0.0) MAT Rieman et al., 1979 0/1 (0.0) PCR Psaroulaki et al., 2014 Little owl Athene noctua http://maps.iucnredlist.org/map.html?id=22689328 Cyprus 0/2 (0.0) PCR Ioannou et al., 2009 Strigidae n.a. n.a. Spain 0/14 (0.0) PCR Astobiza et al., 2011 Northern long-eared 1/4 (25.0) PCR Ioannou et al., 2009 Strigiformes Asio otus http://maps.iucnredlist.org/map.html?id=22689507 Cyprus owl 2/3 (66.6) PCR Psaroulaki et al., 2014 Spotted owlet Athene brama http://maps.iucnredlist.org/map.html?id=22689332 India 1/6 (16.6) CAT Yadav and Sethi 1980 2/5 (40.0) PCR Ioannou et al., 2009 Common barn owl Tyto alba http://maps.iucnredlist.org/map.html?id=22688504 Cyprus Tytonidae 3/5 (60.0) PCR Psaroulaki et al., 2014 n.a. n.a. Spain 0/23 (0.0) PCR Astobiza et al., 2011 0/3 (0.0) PCR Ioannou et al., 2009 Great cormorant Phalacrocorax carbo http://maps.iucnredlist.org/map.html?id=22696792 Cyprus Phalacrocoracidae 0/5 (0.0) PCR Psaroulaki et al., 2014 Suliformes n.a. n.a. Spain 0/2 (0.0) PCR Astobiza et al., 2011 Sulidae n.a. n.a. Spain 0/1 (0.0) PCR Astobiza et al., 2011

Clase Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference Cane toad Rhinella marina http://maps.iucnredlist.org/map.html?id=41065 Fr. Guiana 0/21 (0.0) ELISA Gardon et al., 2001 Bufonidae Toads Bufo spp. India 0/66 (0.0) CAT Yadav and Sethi, 1979 Amphibia Anura Dicroglossidae Rana tigerina Hoplobatrachus tigerinus http://maps.iucnredlist.org/map.html?id=58301 India 0/7 (0.0) CAT Yadav and Sethi, 1979

Leptodactylidae Smoky jungle frog Leptodactylus pentadactylus http://maps.iucnredlist.org/map.html?id=57154 Fr. Guiana 0/20 (0.0) ELISA Gardon et al., 2001

11/20 (55.0) PCR Yadav and Sethi, 1979 Natricidae Snakes Natrix natrix & Naja naja http://maps.iucnredlist.org/map.html?id=14368 India 11/48 (23.0) CAT Yadav and Sethi, 1979

1/2 (50.0) PCR Yadav and Sethi, 1979

2/5 (40.0) CAT Yadav and Sethi, 1979 Squamata Colubridae Chinese rat snake Ptyas korros http://www.catalogueoflife.org/col/details/species India 7/23 (30.4) CAT Yadav and Sethi, 1980

Sauropsida n.a. ISO Yadav and Sethi, 1980

http://maps.iucnredlist.org/map.html?id=178416 1/1 (100.0) PCR Yadav and Sethi, 1979 Varanidae Indian monitor Varanus indicus India n.a. (0.0) CAT Yadav and Sethi, 1980

Pangshura tecta http://maps.iucnredlist.org/map.html?id=46370 India n.a. (0.0) CAT Yadav and Sethi, 1980

Testudines Geoemydidae Tortoise 2/16 (12.5) CAT Yadav and Sethi, 1979 Kachuga spp. http://maps.iucnredlist.org/map.html?id=46370 India 4/7 (57.1) PCR Yadav and Sethi, 1979

Anguilliformes Anguillidae Indian mottled eel Anguilla bengalensis http://maps.iucnredlist.org/map.html?id=61668607 India n.a. (0.0) CAT Yadav and Sethi, 1980

Actinopterygii 0/15 (0.0) CAT Yadav and Sethi, 1979 Cypriniformes Cyprinidae Mrigal Cirrhinus mrigala http://maps.iucnredlist.org/map.html?id=166146 India 2/2 (100.0) PCR Yadav and Sethi, 1979

Scientific Name Host Host scientific name Country Pos/N (Prev) Method Reference

USA n.a. (+) PCR Klyachko et al., 2007 Amblyomma americanum Questing Brazil n.a. (+) PCR Machado-Ferreira et al., 2011 Amblyomma geayi Three toed sloth Bradypus tridactylus French Guiana 14/16 (87.5) PCR Davoust et al., 2014 Python Python sp. Spain 0/1 (0.0) PCR/RLB Toledo et al., 2009 Amblyomma latum Questing Spain 0/1 (0.0) PCR/RLB Toledo et al., 2009 Agile wallaby Macropus agilis Australia 0/6 (0.0) PCR Cooper et al., 2013 Common wallaroo Macropus robustus Australia 0/4 (0.0) PCR Cooper et al., 2013 Amblyomma triguttatum Eastern grey kangaroo Macropus giganteus Australia 14/43 (28.0) PCR Cooper et al., 2013 Kangaroos Macropus rufus, M. giganteus Australia 13/3000 (0.4) ISO Pope et al., 1960 Rufous bettong Aepyprymmus rufescens Australia 0/2 (0.0) PCR Cooper et al., 2013 ISO Rehacek et al., 1991 Slovakia 0/16 (0) PCR Smetanova et al., 2006 Questing Slovakia & Hungary 2/43 (4.6) PCR Spitalska et al., 2003 Spain 18/265 (27.7) PCR/RLB Toledo et al., 2009 Dermacentor marginatus Red deer Cervus elaphus Spain 0/2 (0.0) PCR Astobiza et al., 2011 Wild boar Sus scrofa Spain 0/6 (0.0) PCR Astobiza et al., 2011 Wild mammals (red deer, Cervus elaphus, Sus scrofa, wild boar, red fox, European Vulpes vulpes, Erinaceus Spain 0/23 (0.0) PCR/RLB Toledo et al., 2009 hedgehog, beech marten) europaeus, Martes foina European badger Meles meles Spain 0/3 (0.0) PCR Astobiza et al., 2011 ISO Rehacek et al., 1991 Slovakia Questing 0/9 (0.0) PCR Smetanova et al., 2006 Dermacentor reticulatus Slovakia & Hungary 0/1 (0.0) PCR Spitalska et al., 2003 Red deer Cervus elaphus Spain 0/7 (0.0) PCR Astobiza et al., 2011 Red fox Vulpes vulpes Spain 0/28 (0.0) PCR Astobiza et al., 2011 Wild boar Sus scrofa Spain 0/5 (0.0) PCR Astobiza et al., 2011

Scientific Name Host Name scientific host Country Pos./N (Prev.) Method Reference

Dermacentor spp. Questing Germany 0/666 (0.0) PCR Pluta et al., 2010 Spain n.a. (0.0) PCR Barandika et al., 2008 Questing Slovakia n.a. (+) ISO Rehacek et al., 1991 Haemaphysalis concinna Slovakia & Hungary 0/26 (0.0) PCR Spitalska et al., 2003 Red fox Vulpes vulpes Spain 0/2 (0.0) PCR Astobiza et al., 2011 Lepus europaeus, Oryctolagus European hare & rabbit Spain 0/5 (0.0) PCR/RLB Toledo et al., 2009 Haemaphysalis hispanica cuniculus Questing Spain 0/1 (0.0) PCR/RLB Toledo et al., 2009 Common northern Haemaphysalis humerosa Isoodon macrourus Australia 0/250 (0.0) PCR Cooper et al., 2013 bandicoot Slovakia n.a. (+) ISO Rehacek et al., 1991 Questing Slovakia & Hungary 0/7 (0.0) PCR Spitalska et al., 2003 Haemaphysalis inermis Red deer Cervus elaphus Spain 0/24 (0.0) PCR Astobiza et al., 2011 Roe deer Capreolus capreolus Spain 0/4 (0.0) PCR Astobiza et al., 2011 2/2 (100.0) PCR Ioannou et al., 2011* Cypriot mouflon Ovis orientalis ophion Cyprus 2/2 (100.0) PCR Psaroulaki et al., 2014a Slovakia n.a. (+) ISO Rehacek et al., 1991 Questing n.a. (+) PCR Barandika et al., 2008 Haemaphysalis punctata Spain 0/1 (0.0) PCR/RLB Toledo et al., 2009 Red deer Cervus elaphus Spain 0/1 (0.0) PCR Astobiza et al., 2011 Red fox Vulpes vulpes Spain 0/2 (0.0) PCR Astobiza et al., 2011 Roe deer Capreolus capreolus Spain 0/6 (0.0) PCR Astobiza et al., 2011 5/41 (12.2) PCR Ioannou et al., 2011 Haemaphysalis sulcata Cypriot mouflon Ovis orientalis ophion Cyprus 5/41 (12.2) PCR Psaroulaki et al., 2014a Haemaphysalis sp. House sparrow Passer domesticus France 0/1 (0.0) PCR Socolovschi et al., 2012

Scientific Name Host Name scientific host Country Pos./N (Prev.) Method Reference

2/15 (13.3) PCR Ioannou et al., 2011 Hyalomma excavatum Cypriot mouflon Ovis orientalis ophion Cyprus 2/15 (13.3) PCR Psaroulaki et al., 2014a Birds (Booted eagle, Hieraaetus pennatus, Buteo Spain 0/1 (0.0) PCR/RLB Toledo et al., 2009 common buzzard, red kite) buteo, Milvus milvus Questing Spain 61/701 (8.7) PCR/RLB Toledo et al., 2009 Hyalomma lusitanicum Red deer Cervus elaphus Spain 3/10 (30.0) PCR/RLB Toledo et al., 2009 Wild mammals (wild boar, Sus scrofa, Vulpes vulpes, red fox, European hedgehog, Erinaceus europaeus, Martes Spain 0/11 (0.0) PCR/RLB Toledo et al., 2009 beech marten) foina Birds (Booted eagle, Hieraaetus pennatus, Buteo Spain 0/1 (0.0) PCR/RLB Toledo et al., 2009 common buzzard, red kite) buteo, Milvus milvus Turdus merula, Sylvia Common blackbird, atricapilla, Phoenicurus blackcap, common Redstart, phoenicurus, Saxicola rubetra, whinchat, whitethroat, Italy 10/37 (27.0) PCR Toma et al., 2014 Hyalomma marginatum Sylvia communis, Luscinia nightingale, European megarhynchos, Pernis honey buzzard apivorus 0/1 (0.0) PCR Ioannou et al., 2011 Cypriot mouflon Ovis orientalis ophion Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014a Turdus merula, Sylvia Common blackbird, atricapilla, Phoenicurus blackcap, common Redstart, phoenicurus, Saxicola rubetra, Hyalomma rufipes whinchat, whitethroat, Italy 29/71 (40.0) PCR Toma et al., 2014 Sylvia communis, Luscinia nightingale, European megarhynchos, Pernis honey buzzard apivorus Eurasian reed warbler Acrocephalus scirpaceus France 0/1 (0.0) PCR Socolovschi et al., 2012 Turdus merula, Sylvia Common blackbird, atricapilla, Phoenicurus blackcap, common Redstart, Hyalomma spp. phoenicurus, Saxicola rubetra, whinchat, whitethroat, Italy 29/71 (40.0) PCR Toma et al., 2014 Sylvia communis, Luscinia nightingale, European megarhynchos, Pernis honey buzzard apivorus

Scientific Name Host Name scientific host Country Pos./N (Prev.) Method Reference

Bank vole Myodes glareolus Italy 2/88 (2.2) PCR Pascucci et al., 2015* Ixodes acuminatus Wood mouse Apodemus sylvaticus Italy 4/88 (4.5) PCR Pascucci et al., 2015* European badger Meles meles Spain 0/18 (0.0) PCR Astobiza et al., 2011 Ixodes canisuga Red fox Vulpes vulpes Spain 0/6 (0.0) PCR Astobiza et al., 2011 Ixodes frontalis Song thrush Turdus philomelos Spain 0/2 (0.0) PCR Astobiza et al., 2011 0/3 (0.0) PCR Ioannou et al., 2011 Cypriot mouflon Ovis orientalis ophion Cyprus 0/3 (0.0) PCR Psaroulaki et al., 2014a Ixodes gibossus European hare Lepus europaeus Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014a Red fox Vulpes vulpes indutus Cyprus 0/3 (0.0) PCR Psaroulaki et al., 2014a American mink Neovison vison Spain 0/3 (0.0) PCR Astobiza et al., 2011 European badger Meles meles Spain 0/23 (0.0) PCR Astobiza et al., 2011 Genet Genetta genetta Spain 0/1 (0.0) PCR Astobiza et al., 2011 Ixodes hexagonus Questing Spain n.a. (0.0) PCR Barandika et al., 2008 Red fox Vulpes vulpes Spain 0/7 (0.0) PCR Astobiza et al., 2011 Stone marten Martes foina Spain 0/3(0.0) PCR Astobiza et al., 2011 Weasel Mustela nivalis Spain 0/1 (0.0) PCR Astobiza et al., 2011 Common northern Ixodes holocyclus Isoodon macrourus Australia 10/30 (33.3) PCR Cooper et al., 2013 bandicoot Bank vole Myodes glareolus Italy n.a. (0.0) PCR Pascucci et al., 2015

European badger Meles meles Spain 0/19 (0.0) PCR Astobiza et al., 2011 Ixodes ricinus European hare Lepus europaeus Spain 0/1 (0.0) PCR Astobiza et al., 2011

European pine marten Martes martes Spain 0/1 (0.0) PCR Astobiza et al., 2011

Scientific Name Host Name scientific host Country Pos./N (Prev.) Method Reference

Spain n. a. (0.0) PCR Barandika et al., 2008 Germany 19/1000 (1.9) PCR Hildebrandt et al., 2011 Slovakia n.a. (+) ISO Rehacek et al., 1991 Germany 0/1716 (0.0) PCR Rehacek et al., 1993 Austria 2/298 (0.6) CFT Rehacek et al., 1994 Luxembourg 0/1394 (0.0) PCR Reye et al., 2010 Questing Belarus 5/453 (1.1) PCR Reye et al., 2013 Slovakia 1/327 (0.3) PCR Smetanova et al., 2006 Slovakia & Hungary 4/158 (2.5) PCR Spitalska et al., 2003 Ixodes ricinus Poland 191/1200 (15.9) PCR Szymańska-Czerwińska et al., 2013 Spain 0/8 (0.0) PCR/RLB Toledo et al., 2009 Netherlands 0/1891 (0.0) PCR Sprong et a., 2012 Spain 0/40 (0.0) PCR Astobiza et al., 2011 Red deer Cervus elaphus Netherlands 0/176 (0.0) PCR Sprong et al., 2012 Red fox Vulpes vulpes Spain 0/18 (0.0) PCR Astobiza et al., 2011 Roe deer Capreolus capreolus Spain 0/404 (0.0) PCR Astobiza et al., 2011 Small mammals n.a. Germany 0/892 (0.0) PCR Rehacek et al., 1993 Wood mouse Apodemus spp Italy 4/88 (4.5) PCR Pascucci et al., 2015 Bank vole Myodes glareolus Italy n.a. (0.0) PCR Pascucci et al., 2015 Turdus merula, Sylvia Common blackbird, atricapilla, Phoenicurus blackcap, common redstart, Ixodes spp. phoenicurus, Saxicola rubetra, whinchat, whitethroat, Italy 1/6 (16.6) PCR Toma et al., 2014 Sylvia communis, Luscinia nightingale, European megarhynchos, Pernis honey buzzard apivorus

Scientific Name Host Name scientific host Country Pos./N (Prev.) Method Reference

Eurasian blackbird Turdus merula France 0/4 (0.0) PCR Socolovschi et al., 2012 Eurasian blackcap Sylvia atricapilla France 0/3 (0.0) PCR Socolovschi et al., 2012 European robin Erithacus rubecula France 0/4 (0.0) PCR Socolovschi et al., 2012 Field mouse Apodemus spp. Italy 1/88 (1.1) PCR Pascucci et al., 2015

Ixodes spp. Brünnich’s guillemot Uria lomvia Norway 0/20 (0.0) PCR Duron et al., 2014 King penguin Aptenodytes patagonicus Crozet Archipelago 0/20 (0.0) PCR Duron et al., 2014 Red-faced cormorant Phalacrocorax urile Russia 0/20 (0.0) PCR Duron et al., 2014 Rhinoceros auklet Cerorhinca monocerata Canada 7/14 (50.0) PCR Duron et al., 2014 Tufted puffin, Brünnich’s Fratercula cirrhata, Uria USA 0/20 (0.0) PCR Duron et al., 2014 guillemot lomvia Cyprus 3/15 (20.0) PCR Ioannou et al., 2009 Chukar partridge Alectoris chukar Cyprus 2/2 (100.0) PCR Psaroulaki et al., 2014a Ixodes ventalloi European hare Lepus europaeus Cyprus 5/9 (55.5) PCR Psaroulaki et al., 2014a Red fox Vulpes vulpes Cyprus 0/3 (0.0) PCR Psaroulaki et al., 2014a 14/35 (40.0) PCR Ioannou et al., 2011 Cypriot mouflon Ovis orientalis ophion Cyprus 14/35 (40.0) PCR Psaroulaki et al., 2014a n.a. (0.0) PCR Barandika et al., 2008 Rhipicephalus bursa Questing Spain 0/16 (0.0) PCR/RLB Toledo et al., 2009 Wild boar Sus scrofa Spain 0/2 (0.0) PCR Astobiza et al., 2011 European hare Lepus europaeus Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014a H. pennatus, B. buteo, M. Birds Spain 0/1 (0.0) PCR/RLB Toledo et al., 2009 milvus European badger Meles meles Spain 0/14 (0.0) PCR Astobiza et al., 2011

European hare Lepus europaeus Cyprus 0/3 (0.0) PCR Psaroulaki et al., 2014a Rhipicephalus pusillus European hare & rabbit L. europaeus, O. cuniculus Spain 0/22 (0.0) PCR/RLB Toledo et al., 2009

Questing Spain 1/47 (2.1) PCR/RLB Toledo et al., 2009 C. elaphus, S. scrofa, V. vulpes, Wild mammals Spain 0/28 (0.0) PCR/RLB Toledo et al., 2009 E. europaeus, M. foina

Scientific Name Host Name scientific host Country Pos./N (Prev.) Method Reference

Cypriot mouflon Ovis orientalis ophion Cyprus 2/2 (100.0) PCR Ioannou et al., 2011 House sparrow Passer domesticus France 0/3 (0.0) PCR Socolovschi et al., 2012 European hare & rabbit L. europaeus, O. cuniculus Spain 0/1 (0.0) PCR/RLB Toledo et al., 2009 European hare Lepus europaeus Cyprus 9/14 (64.2) PCR Psaroulaki et al., 2014a

Rhipicephalus sanguineus Questing Spain n.a. (0.0) PCR Barandika et al., 2008 Spain 2/14 (14.3) PCR/RLB Toledo et al., 2009 Red fox Vulpes vulpes Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014a Roe deer Capreolus capreolus Spain 0/1 (0.0) PCR Astobiza et al., 2011 Wild mammals (red deer, C. elaphus, S. scrofa, E. wild boar, European Spain 0/24 (0.0) PCR/RLB Toledo et al., 2009 europaeus, M.es foina hedgehog, beech marten) Bank vole Myodes glareolus Italy n.a. (0.0) PCR Pascucci et al., 2015

Wood mouse Apodemus spp. Italy n.a. (0.0) PCR Pascucci et al., 2015

Cypriot mouflon Ovis orientalis ophion Cyprus 4/10 (40.0) PCR Ioannou et al., 2011 Rhipicephalus turanicus Bank vole Myodes glareolus Italy 4/10 (40.0) PCR Psaroulaki et al., 2014a European hare Lepus europaeus Cyprus 10/32 (31.0) PCR Psaroulaki et al., 2014a Red fox Vulpes vulpes indutus Cyprus 3/19 (15.7) PCR Psaroulaki et al., 2014a Ticks (n.a.) Poland n.a. (+) ISO Tylewska-Wierzbanowska et al., 1991

Scientific Name Host Name scientific host Country Pos./N (Prev.) Method Reference

Argas reflexus Pigeon tower Columba sp. France 6/20 (30.0) PCR Stein et al., 1999 Carios capensis Brown pelican Pelecanus occidentalis USA 64/64 (100.0) PCR Reeves et al., 2006 Cape Verde shearwater, Calonectris edwardsii, Sula Cape Verde 16/16 (100.0) PCR Duron et al., 2014 brown booby leucogaster Humboldt penguin Spheniscus humboldti Chile 3/3 (100.0) PCR Duron et al., 2014 Peruvian pelican, Peruvian Pelecanus thagus, Sula Peru 5/5 (100.0) PCR Duron et al., 2014 Ornithodoros capensis s.l. booby variegata Sooty tern Onychoprion fuscatus Mozambique 28/28 (100.0) PCR Duron et al., 2014 Spain 20/20 (100.0) PCR Duron et al., 2014 Yellow-legged gull Larus michahellis Tunisia 20/20 (100.0) PCR Duron et al., 2014

Ctenocephalides canis European hare Lepus europaeus Cyprus 0/2 (0.0) PCR Psaroulaki et al., 2014b*

Rats Rattus sp. Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014b

Red fox Vulpes vulpes Cyprus 8/18 (44.4) PCR Psaroulaki et al., 2014b European hare Lepus europaeus Cyprus 0/4 (0.0) PCR Psaroulaki et al., 2014b Ctenocephalides felis Rats Rattus sp. Cyprus 6/41 (14.6) PCR Psaroulaki et al., 2014b Red fox Vulpes vulpes Cyprus 2/3 (66.6) PCR Psaroulaki et al., 2014b European hare Lepus europaeus Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014b Xenopsylla cheopis Rats Rattus sp. Cyprus 9/83 (10.3) PCR Psaroulaki et al., 2014b

Order Family Common name Scientific name Country Origin Pos/N (Prev.) Method Reference

Arab Emirates C 7/170 (4.1) ELISA Chaver et al., 2012 Jordan C 0/10 (0.0) CFT Greth at al., 1992 Barhain C 0/2 (0.0) CFT Greth at al., 1992 Arabian Oryx Oryx leucoryx USA C 0/8 (0.0) CFT Greth at al., 1992 Qatar C 1/2 (50.0) CFT Greth at al., 1992 C 18/189 (9.5) CFT Greth at al., 1992 Saudi Arabia C 45/96 (46.9) ELISA Hussein et al., 2012 Black buck Antilope cervicapra Arab Emirates C 2/36 (5.5) ELISA Chaver et al., 2012 Blackfaced impala Aepyceros melampus petersi Portugal Z 0/1 (0.0) PCR Cumbassá et al., 2015 Dama gazelle Nanger dama Arab Emirates C Patology Lloyd et al., 2010 Dorcas gazelle Gazella dorcas Arab Emirates C 1/3 (33.3) ELISA Chaver et al., 2012 Grant´s gazelle Nanger granti Arab Emirates C 1/15 (6.6) ELISA Chaver et al., 2012 Artiodactyla Bovidae Impala Aepyceros melampus Arab Emirates C 0/7 (0.0) ELISA Chaver et al., 2012 Kafue lechwe Kobus leche kafuensis Zambia F + Krauss et al., 1986 Laristan sheep Ovis laristanicus Arab Emirates C 0/3 (0.0) ELISA Chaver et al., 2012 Lessur kudu Tragelaphus imberbis Arab Emirates C 1/20 (5.0) ELISA Chaver et al., 2012 Mouflon Ovis orientalis musimon Slovakia Z 1/4 (25.0) ELISA Dorko et al., 2009 Arab Emirates C 0/3 (0.0) ELISA Chaver et al., 2012 Mountain gazelle Gazella gazella Saudi Arabia C 17/232 (7.3) ELISA Hussein et al., 2012 Muskox Ovibos moschatus Germany Z + CFT Schroder et al., 1998 Roan antelope Hippotragus niger niger Portugal Z + PCR Clemente et al., 2008 Arab Emirates C 2/6 (33.3) ELISA Chaver et al., 2012 Sand gazelle Gazella leptoceros Saudi Arabia C 37/227 (18.3) ELISA Hussein et al., 2012 Speke´s gazelle Gazella spekei Arab Emirates C 6/70 (8.6) ELISA Chaver et al., 2012 Watebuck Kobus ellipsiprymnus Portugal Z + PCR Clemente et al., 2008

Order Family Common name Scientific name Country Origin Pos/N (Prev.) Method Reference

12/1012 (1.2) SERO Galiero et al., 1996 Water buffalo Bubalus bubalis Italy F Bovidae 14/164 (17.5) PCR Perugini et al., 2009

White antelope Addax nasomaculatus Portugal Z 2/15 (13.3) PCR Cumbassá et al., 2015

Slovakia Z 20/60 (33.3) ELISA Dorko et al., 2009 Fallow deer Dama dama Germany F + CFT/ELISA Simmert et al., 1998 Korea C 0/30 (0.0) ELISA Jang et al., 2011 Sika deer Cervus nippon China F 166_1347 (12.3) ELISA Cong et al., 2015 Pampas deer Ozotoceros bezoarticus Uruguay E 5/22 (22.0) ELISA Hernández et al., 2007 Artiodactyla Portugal Z 0/1 (0.0) PCR Cumbassá et al., 2015 F 6/23 (26.1) PCR González-Barrio et al., 2015 Cervidae F 9/25 (36.0) ELISA González-Barrio et al., 2015 Red deer Cervus elaphus F 1/29 (3.4) PCR González-Barrio et al., 2015 Spain F 173/600 (28.8) ELISA González-Barrio et al., 2015 F 32/80 (40.0) IFA Ruiz-Fons et al., 2008 F 4/32 (12.5) PCR Ruiz-Fons et al., 2008 Germany Z + n.a. Gaukler and Krauss, 1974 Wapiti Cervus canadensis Korea F 10/604 (1.7) ELISA Jang et al., 2011 1/8 (12.5) PCR Laricchiuta et al., 2009 Felidae Lion Panthera leo Italy S 2/8 (25.0) IFA Laricchiuta et al., 2009 Carnivora n.a. (10.0) IFA Torina et al., 2007 Ursidae Brown bear Ursus arctos Croatia C 2/9 (22.2) CFT Madic et al., 1993 Cetartiodactyla Giraffidae Giraffe Giraffa camelopardalis Portugal Z 1/1 (100.0) PCR Cumbassá et al., 2015

Order Family Common name Scientific name Country Origin Pos/N (Prev.) Method Reference

2/97 (2.0) MAT Kocianova et al., 1993

Columbiformes Columbidae Rock dove Columba livia Slovakia SD 12/67 (17.9) MAT Kocianova et al., 1993

16/97 (16.5) MAT Kocianova et al., 1993

+ MAT Schatmz et al., 1977 Galliformes Phasianidae Common quail Coturnix coturnix Germany E n.a. ISO Schatmz et al., 1977 Lagomorpha Leporidae European rabbit Oryctolagus cuniculus Spain F 9/108 (8.3) ELISA González-Barrio et al., 2015 n.a. n.a. Marine mammals n.a. Germany Z + n.a. Jurczynski et al., 2005 Perissodactyla Equidae Plains zebra Equus quagga Portugal Z 0/2 (0.0) PCR Cumbassá et al., 2015 Perissodactyla Rhinocerotidae White rhinoceros Ceratotherium simum Portugal Z 0/1 (0.0) PCR Cumbassá et al., 2015 Ramphastidae Toucan Ramphastos toco USA C Pathology PCR Shivaprarad et al., 2008 Piciformes Cacatuidae Cockatiel Nymphicus hollandicus USA C Pathology PCR Shivaprarad et al., 2008 Psittacidae Canary-winged parakeet Brotogeris chiriri USA C Pathology PCR Shivaprarad et al., 2008 Hawk-headed parrot Deroptyus accipitrinus USA C Pathology PCR Shivaprarad et al., 2008 Psittaciformes Red-fronted parakeet Cyanoramphus novaezelandiae USA C Pathology PCR Shivaprarad et al., 2008 Psittaculidae Golden mantle rosella Platycercus eximius USA C Pathology PCR Shivaprarad et al., 2008 Swainson's Blue Mountain rainbow lorikeets Trichoglossus haematodus moluccanus USA Z Pathology n.a. Woc-Colburn et a.l, 2008

Objetivos

El Objetivo Principal de esta Tesis Doctoral es estimar cuál es el papel que juega la fauna silvestre ibérica en la ecología de Coxiella burnetii y evaluar potenciales métodos para su control en poblaciones de fauna silvestre.

Los Objetivos Específicos para abordar el objetivo general son:

1.- Determinar el papel como reservorio de C. burnetii de varias especies de fauna silvestre, en concreto del ciervo rojo y del conejo de monte; así como el estado de C. burnetii en sus poblaciones y los factores de riesgo que modulan la exposición al patógeno.

2.- Evaluar la dinámica de exposición a C. burnetii en el tiempo en una población de ciervo rojo en la que C. burnetii es endémica y caracterizar la epidemiología del patógeno en este tipo de escenarios.

3.- Caracterizar los factores que modulan en el tiempo la exposición a C. burnetii en ciervo rojo.

4.- Caracterizar los genotipos de C. burnetii que infectan a la fauna silvestre y compararlos con aquellos descritos en ganado doméstico, garrapatas y casos clínicos de fiebre Q humanos.

5.- Determinar cuáles son las vías de excreción de C. burnetii en ciervo rojo, conejo de monte y jabalí.

6.- Diseñar y evaluar la eficacia de programas de vacunación con vacunas inactivadas de fase I como método de control en poblaciones de ciervo rojo.

Capítulo II. Epidemiología

de Coxiella burnetii en

fauna silvestre ibérica

Capítulo II. 1

100

Estado de Coxiella burnetii en las poblaciones de ciervo rojo (Cervus

elaphus) en la península ibérica y factores de riesgo asociados

Host and Environmental Factors Modulate the Exposure of Free-Ranging and

Farmed Red Deer (Cervus elaphus) to Coxiella burnetii

David González-Barrio, Ana Luisa Velasco Ávila, Mariana Boadella, Beatriz Beltrán-

Beck, José Ángel Barasona, João P. V. Santos, João Queirós, Ana L. García-Pérez,

Marta Barral, Francisco Ruiz-Fons

Applied and Environmental Microbiology. 2015. 81 (18): 6223-6231

101

Resumen

El control de patógenos multihospedadores como Coxiella burnetii debe contar con la información precisa sobre el papel desempeñado por los principales reservorios. El objetivo de este trabajo fue determinar la implicación del ciervo rojo (Cervus elaphus) en la ecología de Coxiella burnetii. Suponiendo que las poblaciones de ciervos de amplias zonas geográficas dentro de un contexto europeo estarían expuestos a Coxiella burnetii, y por lo tanto, formulamos la hipótesis de que una serie de factores podría modular esta la exposición de los ciervos a Coxiella burnetii. Para testar esta hipótesis, diseñamos un estudio retrospectivo de 47 poblaciones de ciervo en la Peninsula Ibérica, en las cuales

1751 sueros y 489 muestras de bazos fueron tomados. Los sueros fueron analizados por ensayo por inmunoadsorción ligado a enzimas (ELISA) con el fin de estimar la exposición a Coxiella burnetii, y las muestras de bazo fueron analizadas por medio de PCR con el fin de estimar la prevalencia de infección sistémica. A continuación, reunimos 23 variables - dentro de factores ambientales, factores propios del hospedador y factores de manejo – potencialmente moduladores del riesgo de exposición del ciervo a Coxiella burnetii, y se realizaron análisis estadísticos multivariados para identificar los principales factores de riesgo. Veintitres poblaciones fueros seropositivas (48,9%), y el ADN de

Coxiella burnetii en bazo fue detectado en el 50% de las poblaciones analizadas. El análisis estadístico refleja la complejidad de la ecología de Coxiella burnetii y sugiere que a pesar de que el ciervo puede mantener la circulación de C. burnetii sin terceras especies, probablemente se incluyan otras especies de reservorios silvestres y domésticos en el ciclo de vida de Coxiella burnetii.

102

Abstract

The control of multihost pathogens, such as Coxiella burnetii, should rely on accurate information about the roles played by the main hosts. We aimed to determine the involvement of the red deer (Cervus elaphus) in the ecology of C. burnetii. We predicted that red deer populations from broad geographic areas within a European context would be exposed to C. burnetii, and therefore, we hypothesized that a series of factors would modulate the exposure of red deer to C. burnetii. To test this hypothesis, we designed a retrospective survey of 47 Iberian red deer populations from which 1,751 serum samples and 489 spleen samples were collected. Sera were analyzed by enzyme-linked immunosorbent assays (ELISA) in order to estimate exposure to C. burnetii, and spleen samples were analyzed by PCR in order to estimate the prevalence of systemic infections.

Thereafter, we gathered 23 variables— within environmental, host, and management factors—potentially modulating the risk of exposure of deer to C. burnetii, and we performed multivariate statistical analyses to identify the main risk factors. Twenty-three populations were seropositive (48.9%), and C. burnetii DNA in the spleen was detected in 50% of the populations analyzed. The statistical analyses reflect the complexity of C. burnetii ecology and suggest that although red deer may maintain the circulation of C. burnetii without third species, the most frequent scenario probably includes other wild and domestic host species. These findings, taken together with previous evidence of C. burnetii shedding by naturally infected red deer, point at this wild ungulate as a true reservoir for C. burnetii and an important node in the life cycle of C. burnetii, at least in the Iberian Peninsula.

103

Introduction

Coxiella burnetii is a Gram-negative intracellular bacterium that causes Q fever, a disease that affects both humans and animals. Whereas the epidemiological status of C. burnetii in European domestic ruminants is well known (Angelakis & Raoult, 2010), information for wildlife is mostly local and scattered (EFSA, 2010; Ruiz-Fons, 2012). Although the majority of human Q fever outbreaks are linked to the transmission of C. burnetii from domestic ruminants (Roest et al., 2011a; Georgiev et al., 2013), the ability of C. burnetii to infect wild hosts (Ruiz-Fons, 2012; Badubieri, 1959) and its high environmental resistance (Angelakis & Raoult, 2010) make wildlife species potential reservoirs of C. burnetii. Based on this hypothesis, wildlife could maintain C. burnetii and transmit it to wildlife (González-Barrio et al., 2015c), domestic animals (Jado et al., 2012), or humans

(Tozer et al., 2014). It is therefore of paramount relevance (i) to identify those potential wild reservoir species that could, through direct and indirect interactions, transmit C. burnetii to target species (domestic animals and humans) and (ii) to determine which environmental factors are the main drivers of C. burnetii within the most relevant wild reservoirs. Efficient prevention of C. burnetii transmission at the wildlife– domestic- animal–human interface can be approached only once the main reservoirs have been identified and the driving risk factors are known (Viana et al., 2014).

Several wild ruminant species are present and well distributed in Europe; on the premise that they are susceptible to infection by C. burnetii, these could constitute important wild reservoirs of C. burnetii. However, among European wild ruminants, the red deer (Cervus elaphus) could perhaps constitute a potential wild reservoir for C. burnetii due to its geographic distribution, demographic status, importance as game, and behavior. The red deer displays broad global (Flueck et al., 2003; Ludt et al., 2004) and European (Zachos

& Hartl, 2011) geographic distribution, with trends to increasing distribution and density

104

(Acevedo et al., 2008; Apollonio et al., 2010). It is currently one of the most important game species among European large mammals (Milner et al., 2007). Many red deer populations in Europe are subjected to management for hunting (Vicente et al., 2006), and red deer farming has expanded in recent decades as a consequence of the demand for venison and live individuals for population-restocking programs (Hoffman & Wiklund,

2006). Additionally, the gregarious behavior of the red deer (Clutton-Brock et al., 1982;

Vander Wal et al., 2013) promotes the aggregation of individuals. In domestic animals, host density and aggregation are important drivers of C. burnetii transmission (Álvarez et al., 2012; Piñero et al., 2014), and some Iberian red deer populations reach densities higher than 70 deer/km2 (Acevedo et al., 2008). Increasing red deer densities, deer management (including artificial feeding), and gregarious behavior constitute the main factors favoring the transmission of circulating pathogens in red deer populations (Ruiz-

Fons et al., 2008a; Boadella et al., 2010).

Taken together, distribution, demography, management, and behavior point at red deer as one of the most concerning reservoirs of shared pathogens among European wild ruminants; e.g., 44% of red deer in Italy were found to be infected by piroplasms (Zanet et al., 2014), and 60% of Slovakian red deer carried Anaplasma spp. (Vichová et al.,

2014). Therefore, we predicted that C. burnetii would be circulating in red deer populations in Iberia, and we hypothesized that particular environmental, management, and host factors would contribute to the exposure of red deer to C. burnetii. To test these hypotheses, we designed a retrospective epidemiological survey targeting Iberian

(Spanish and Portuguese) red deer populations within their geographic distribution range.

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Figure 1. Spatial distribution of Coxiella burnetii seroprevalence in Iberian red deer and presence of C. burnetii DNA in spleen samples. Each dot represents a surveyed red deer population. Current geographic distribution of the red deer in the Iberian Peninsula is shown in pale orange (Salazar, 2009; Palomo et al., 2007). The number of sera analyzed per population is displayed in numbers. A red asterisk within the sampling size indicates red deer farms. The map of Spain has been split according to bioregions established in the current Spanish wildlife disease surveillance program (Muñoz et al., 2010).

Materials and methods

Survey design. Sera from 47 red deer populations were collected from 2000 to 2012 in mainland Spain and Portugal (Fig. 1). Study populations were selected on the basis of (i) management systems, including unmanaged, naturally free-ranging populations (in game

106 reserves and natural and national parks), managed free-ranging populations, and farms,

(ii) geographic location (location within the different bioregions established for wildlife disease surveillance schemes in mainland Spain (Muñoz et al., 2010) and from different regions in mainland Portugal), and (iii) the range of geographic distribution of red deer

(Fig. 1), in order to obtain spatial representativeness.

Serological analyses. The presence of specific antibodies against C. burnetii phase I and

II antigens in deer sera was analyzed with a comercial indirect enzyme-linked immunosorbent assay (ELISA) (LSIVet Ruminant Q Fever Serum/Milk ELISA kit; Life

Technologies, USA) with an in-house modification in the secondary antibody (protein G- horseradish peroxidase; Sigma-Aldrich, USA) (Stöbel et al., 2002) that was previously validated for wild and domestic ungulates (Ruiz-Fons et al., 2010a). Briefly, for validation, we employed positive (n=8) and negative (n=6) red and roe deer sera analyzed by indirect immunofluorescence assay (IFA), as well as ELISA-positive, PCR-positive and ELISA-negative, PCR-negative cattle (n, 14 and 12, respectively) and sheep (n, 16 and 17, respectively) sera. For each sample, the sample-to-positive-control (SP) ratio was calculated as ((ODs-ODnc)/(ODpc-ODnc))*100, where ODs is the optical density of the sample as measured using a dual-wavelength spectrophotometer (first at 450 nm and then at 620 nm), ODnc is the optical density of the negative control, and ODpc is the optical density of the positive control. All SP values of 40% were considered negative, whereas

SP values of 40% were considered positive.

PCR analyses. Spleen samples were collected from a subset of the populations studied during necropsies performed on hunter-harvested or euthanized farmed deer. Spleen samples from seropositive and seronegative deer were selected for PCR analyses. Total

DNA from spleen samples was purified with the DNeasy blood and tissue kit (Qiagen,

Germany) according to the manufacturer’s protocol

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(http://mvz.berkeley.edu/egl/inserts/DNeasy_Blood_&_Tissue_Handbook.pdf). The

DNA concentration in aliquots was quantified (NanoDrop 2000c/2000 spectrophotometer; Thermo Scientific, USA), and aliquots were frozen at -20°C until the

PCR was performed. Sample cross-contamination during DNA extraction was excluded by including negative controls (nuclease-free water; Promega, USA) that were also tested by PCR. DNA samples were analyzed by a quantitative real-time PCR (qPCR) targeting a transposon-like repetitive region of C. burnetii as described previously (Table 1)

(Tilburg et al., 2010). SsoAdvanced universal probes supermix (Bio- Rad, USA) was used in qPCR according to the specifications of the manufacturers. DNA extraction and PCR were performed in separate laboratorios under biosafety level II conditions (Bio II A cabinet; Telstar, Spain) to avoid cross-contamination. As a positive control in this real- time PCR, we used a DNA extract of Coxiella burnetii from the Coxevac vaccine (CEVA

Santé Animale, France). We considered a sample to be positive at a threshold cycle (CT) value below 40 (Tilburg et al., 2010).

Table 1. Primers and probe used in the qPCR.

a Location in positions of the whole genome sequence of C. burnetii RSA493 (Gen Bank accession number AE016828), encoding the transposase gene of the C. burnetii-specific IS1111a insertion element.

Risk predictor variables. In order to identify factors modulating the risk of exposure of individual red deer to C. burnetii infection, a set of abiotic and biotic variables within three main factors- environment, management, and host (Table 2) -were gathered on the basis of their potential impact on C. burnetii ecology.

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(i) Environmental factors. Both spatial and meteorology-related variables were considered for risk factor modeling. Longitude (X) and latitude (Y) were considered as spatial factors to control for any potential spatial autocorrelation of data. Coordinates were recorded at the sampling-site level with portable global-positioning-system (GPS) devices (Garmin Ltd., Cayman Islands), so all deer from a sampling site were assigned the same X and Y values. The average spring temperature (AvSpT) and the season (Se) in which deer were surveyed (4 categorical classes: spring [Sp], April to June; summer

[Su], July to September; autumn [Au], October to December; and winter [Wi], January to

March) were considered as meteorology- related variables. AvSpT was considered as a potential proxy for C. burnetii environmental survival and as a potential driver of airborne transmission of C. burnetii—probably dependent on air moisture, which is highly correlated with temperature (Ruiz-Fons et al., 2010b) — in the expected shedding season.

The prevalence of C. burnetii shedding is expected to be higher in the spring, when calving takes place, as a recent study suggests (González-Barrio et al., 2014). The season was considered as a proxy of potential year-round variability in infection risk because of the expected predominance of C. burnetii shedding in the spring.

(ii) Management factors. Human interference in deer ecology and behavior was considered on the basis of deer population management systems: (i) unmanaged free- ranging deer populations (Um), (ii) freeranging deer populations managed for hunting purposes (Mg) (high-wire fencing restriction and year-round supplementary feeding), and

(iii) farmed deer populations (Fd) (extensively produced red deer in 6- to 10-ha enclosures).

(iii) Host factors. Different host population and individual host variables were considered, because C. burnetii is a multihost pathogen (Maurin & Raoult, 1999):

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Table 2. Set of variables gathered for risk factor modeling of deer individual exposure to Coxiella burnetii.

Factor Variable Variable description (measure unit) codea Environment X* Longitude (m) Y* Latitude (m) Se* Season (Sp: spring; Su: summer; Au: autumn; Wi: winter) AvSpT* Average spring temperature (ºC)

Managementa EsT*b Estate type (Um: unmanaged free-ranging; Mg: managed free-ranging; Fd: Farmed) Host CFd* Density of cattle farms in the municipality (farms/Km2) 2 SFd Density of sheep farms in the municipality (farms/Km ) GFd Density of goat farms in the municipality (farms/Km2) SrFd* Density of small ruminant farms in the municipality (farms/Km2) RuFd Density of ruminant farms in the municipality 2 (farms/Km ) Cd Density of cattle in the municipality (Animals/Km2) Sd Density of sheep in the municipality (Animals/Km2) Gd Density of goats in the municipality (Animals/Km2) Srd Density of small ruminants in the municipality (Animals/Km2) Rud* Density of ruminants in the municipality (Animals/Km2) RdFi* Environmental favourability for red deer RoFi Environmental favourability for roe deer WbFi Environmental favourability for wild boar HUd* Density of humans in the municipality (people/Km2) HsDi Distance to the nearest human settlement (Km) Sx* Sex (M: male; F: female) Ag* Age class (Cf: calf; Yr: yearling; Sa: sub-adult; Ad: adult) Unclassified Sy* Sampling year a Variables included in the statistical modeling process are marked with an asterisk b This variable was included only in overall (unmanaged plus manage plus farm) deer data set

(a) Densities of domestic ruminants and domestic ruminant farms in the municipality

to which individual deer belong. Domestic ruminant density (densities of cattle

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[Cd], sheep [Sd], and goats [Gd], and of combinations of these ruminants [Rud])

and farm density (CFd, SFd, GFd, small-ruminant farm density [SrFd], and RuFd)

values at the municipality level were calculated on the basis of livestock census

data gathered by the Spanish and Portuguese National Statistics Institutes

(http://www.ine.es and http: //www.ine.pt, respectively) in 2009.

(b) Environmental favorability index. The environmental favorability index (Fi)

ranged from 0 (minimum favorability) to 1 (maximum favorability) for red deer

(RdFi), roe deer (RoFi), and wild boar (WbFi) at Universal Transverse Mercator

(UTM) 10- by 10-km resolution squares, calculated for peninsular Spain

(Acevedo et al., 2010b). This index is a measure of the suitability of a land surface

for a particular species and is well correlated with the real abundance of the

species (Real et al., 2009). Environmental favorability índices of wild ungulates

have not been estimated for Portugal. Therefore, Portuguese populations that were

close to the Spanish border (n=6) (Fig. 1) were linked to the favorability indices

of the closest Spanish UTM 10- by 10-km square. The only population surveyed

in central Portugal could not be associated with any wild ungulate favorability

index and was not considered for risk factor analyses. Red deer, roe deer, and wild

boar have been found to be infected by C. burnetii previously (Astobiza et al.,

2011a; Rijks et al., 2011; Ejercito et al., 1993). No favorability indices for any

other potential wild host of C. burnetii are available for the study area.

were obtained from the Spanish and Portuguese National Statistics Institutes in

2011 and 2010, respectively.

(d) Straight-line Euclidean distance to the nearest human settlement (HsDi). The

distance to the nearest human settlement (town or city) was measured with

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Geographic Information Systems (Quantum GIS; http: //www.qgis.org/es/site/).

Human and their pets may be hosts for C. burnetii and may potentially modulate

the risk of exposure of deer (Maurin & Raoult, 1999). For this reason, HUd and

HsDi were considered for modeling analyses.

(e) Host sex (Sx; male [M] versus female [F]). Among farmed deer, the number of

stags reared was significantly lower than the number of females reared, and

therefore, there was a sex bias in the sample.

(f) Host age (Ag). For free-ranging deer, tooth eruption patterns (Saenz de Buruaga

et al., 1991) were used to estimate the ages of animals 2 years old, whereas for

animals 2 years old, age was determined by the number of cementum annuli of

the incisor 1 root (Hamlin et al., 2000). Farm keepers provided the year of birth

for farmed deer. For analytical purposes, four age classes were established: calf

(Cf; 0 to 1 years old), yearling (Yr; 1 to 2 years old), subadult (Sa; 2 to 3 years

old), and adult (Ad; 3 years old). In free-ranging populations, a conscious negative

bias against calves existed according to reported agerelated C. burnetii

seroprevalence patterns (González-Barrio et al., 2014; Ruiz-Fons et al., 2010c).

Table 3. Average individual seroprevalence number of positive samples over sampling size and associated 95% confidence interval throughout each sampling bioregiona and deer management system. a See Muñoz et al., 2010. bPos, number of psitive samples; n, total number of sample. cNA, not applicable.

Seroprevalence (%) (Pos/N)b (95%CI) Country or bioregion All deer populations Unmanaged deer Managed deer Farmed deer Spain Bioregion 1 4.3% (7/161; 1.8-8.8) 4.3% (7/161; 1.8-8.8) NAc NA Bioregion 2 5.7% (10/174; 2.8-10.3) 14.3% (8/56; 6.4-26.2) 0.0% (0/59; 0.0-6.1) 3.4% (2/59; 0.4-11.7) Bioregion 3 2.7% (18/675; 1.6-4.2) 3.8% (14/372; 2.1-6.2) 1.5% (4/264; 0.4-3.8) 0.0% (0/39; 0.0-9.0) Bioregion 4 1.7% (2/116; 0.2-6.1) 1.3% (1/79; 0.0-6.7) 0.0% (0/6; 0.0-45.9) 3.2% (1/31; 0.1-16.7) Bioregion 5 34.6% (175/506; 30.4-38.9) 14.3% (5/35; 4.8-30.2) NA 36.1% (170/471; 31.7-40.6) Portugal 1.7% (2/119; 0.2-5.9) 1.7% (2/119; 0.2-5.9) NA NA Total 12.2% (214/1751; 10.7-13.9) 4.5% (37/822; 3.2-6.2) 1.2% (4/329) 0.00-0.03 28.8% (173/600; 25.2-32.6)

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The year in which each individual deer was sampled (Sy) was additionally considered as a survey-associated factor modulating the risk of exposure of deer to C. burnetii (Piñero et al., 2014).

Statistical analyses. Four different data sets were employed to test for the main hypothesis of this study -the modulating effect of risk factors on the exposure of deer to

C. burnetii- in order to seek major driving factors, including or not including the management system (an expected major epidemiological driver according to existing literature on wild ungulate pathogen dynamics). Data sets included (i) overall deer populations studied, (ii) unmanaged free-roaming deer populations, (iii) managed freeroaming deer populations, and (iv) deer farms. The deer management system was included in modeling of the data set that included all deer in order to test for the effect of management on the risk of exposure of deer to C. burnetii. Within each data set and with the aim of reducing the interference of multicollinearity among predictor variables in modeling output, a correlation matrix (Spearman’s rank tests) of continuous variables was built. Therefore, only uncorrelated variables (Spearman’s rho, 0.4) were selected for statistical modeling (Table 2).

For risk factor modeling, multivariate logistic regression models— generalized linear mixed models (McCulloch et al., 2008) fitted with a binomial distribution and a logit link function—were built (lme4 package for R) to test the influence of different potential risk factors (Table 2) on the risk of exposure of individual deer to C. burnetii. The individual status of anti-Coxiella burnetii antibodies was entered as a response variable (coded as 0 for an animal testing negative and as 1 for an animal testing positive) in the model. The location of origin of deer was entered as a random variable in the modeling process.

Models were built by following a forward stepwise procedure with the aim of identifying the main modulating factors of the exposure of deer to C. burnetii. The Akaike

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information criterion (AIC) and the AIC increment (AIC) were considered in order to

select the best-fitted model (i.e., with the lowest AIC value and with a AIC of 2 (Burham

& Anderson, 2002)). The statistical uncertainty associated with the estimation of

individual prevalence values was assessed by calculating the associated Clopper- Pearson

exact 95% confidence interval (95% CI).

Results

A total of 1,751 serum samples were analyzed: 822 (46.9%) from unmanaged populations

(n=27), 329 (18.8%) from managed populations (n=14), and 600 (34.3%) from farmed

populations (n=6). Of the 1,629 samples for which sex could be recorded, 1,147 (70.4%)

were from females and 482 (29.6%) were from males. For 1,593 samples, age could be

recorded; 100 samples (6.3%) belonged to calves, 240 (15.1%) belonged to yearlings, 251

(15.7%) belonged to subadults, and 1,002 (62.9%) belonged to adults. Age and sex could

be recorded for 1,560 individuals at the time. Average individual seroprevalences by

bioregion and deer management system are shown in Table 3.

Table 4. Average seroprevalence values, numbers of positive samples over sampling size, and associated exact 95% confidence interval by deer sex and age.aPos, number of postive samples; N, total number of samples.

Seroprevalence (%) Sex Age class (Pos/N)a (95%CI) Male Calf 2.7% (1/37; 0.1-14.2) Yearling 1.6% (1/61; 0.0-8.8) Sub-adult 3.1% (1/32; 0.1-16.2) Adult 3.9% (13/336; 2.1-6.5) Subtotal, male 3.5% (17/482; 2.1-5.6) Female Calf 2.6% (1/38; 0.1-13.8) Yearling 9.0% (16/177; 5.3-14.3) Sub-adult 33.0% (72/218; 26.8-39.7) Adult 15.4% (102/661; 12.8-18.4)

Subtotal, female 16.9% (194/1,147; 14.8-19.2)

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All IFA-positive red and roe deer sera presented SP values of 100, whereas IFA-negative

sera had SP values of ˂25 (SP cutoff for positivity, 40). ELISA-positive, PCR-positive

cattle and sheep sera displayed SP values of ˃70 and ˃100, respectively, whereas ELISA-

negative, PCR-negative cattle and sheep sera had SP values of ˂30 (Ruiz-Fons et al.,

2010a). Therefore, with the controls employed in our validation approach, the ELISA

reached 100% sensitivity and specificity for a positive cutoff SP of ˃40. Twenty-three of

the 47 deer populations surveyed (48.9%) had at least one seropositive sample; Four out

of six deer farms (66.7%) and 55.6% of unmanaged free-ranging populations (15/ 27) had

seropositive animals, in contrast to 21.4% of managed free-ranging deer populations

(3/14). Seven of the 47 red deer populations (14.9%) had average individual

seroprevalences of10%. Average seroprevalence values by sex and age are shown in

Table 4.

Table 5. Best-fit model output throughout the deer data seta. a The statistc (Z), the coeficient (β),

its associated standard error (SE), the significance value (P), the model Akaike information criterion (AIC), the AIC increment (ΔAIC), and the explained deviance (ED) are shown. b c Abbreviations of variables are explained in Table 2. NS, P>0.05; *, P <0.05; **, P <0.01; ***, P <0.001

Data set Variableb Z β SE Pc AIC ΔAIC ED (%) Unmanaged + Managed + Intercept -2.561 -3.745 1.463 * Farmed HUd 2.707 0.019 0.007 ** Se -2.476 -0.690 0.279 * 364.105 19.206 7.172 AvSpT 2.008 0.204 0.101 * Rud -1.573 -0.017 0.011 NS Unmanaged Intercept -1.331 -1932 1.473 NS HUd 1.788 0.053 0.030 NS 220.417 2.525 3.723 Se -1.587 -0.546 0.344 NS

RdFi -0.862 -1.193 1.382 NS Managed Intercept -2.429 -20.233 8.331 * 39.971 4.741 16.557 AvSpT 2.042 1.134 0556 * Farmed Intercept -2.926 -2.119 0.724 ** Se 3.705 1.983 0.535 *** 91.102 16.224 19.572 RuD -3.499 -0.186 0.053 ***

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A total of 489 spleen samples were analyzed by qPCR (Fig. 1); 305, 155, and 29 spleen samples came from unmanaged, managed, and farmed deer populations, respectively.

Among all spleen samples, 5.7% (28/489) (95% CI, 3.8 to 8.2%) were qPCR positive

(cycle threshold range for positive samples, 32.1 to 39.9). The prevalences of C. burnetii

DNA in spleen were 6.2% (19/305) (95% CI, 3.8 to 9.6%), 5.2% (8/155) (95% CI, 2.3 to

9.9%), and 3.5% (1/29) (95% CI, 0.1 to 17.8%) in unmanaged, managed, and farmed deer populations, respectively. Ten of 140 males (7.1%; 95% CI, 3.5 to 12.6%) and 18 of 234 females (7.7%; 95% CI, 4.6 to 11.9%) were qPCR positive. One of 10 calves analyzed

(10.0%; 95% CI, 0.3 to 44.5%), 5 of 41 juveniles (12.2%; 95% CI, 4.1 to 26.2%), 2 of 19 subadults (10.5%; 95% CI, 1.3 to 33.1%), and 20 of 302 adults (6.6%; 95% CI, 4.1 to

10.1%) were positive for C. burnetii DNA in the spleen by qPCR.

Twenty-six deer populations were studied for the prevalence of C. burnetii DNA in spleen samples. Of these, 12 were seronegative and 14 had at least one seropositive individual.

Thirteen of those 26 deer populations (50.0%) had at least one positive spleen sample; 8 were seropositive and 5 were seronegative.

The best-fitted general model for risk factors for the exposure of deer to C. burnetii (Table

5) retained variables within the host and environment factors. Human density and the average spring temperature were positively related to increasing risks of exposure to C. burnetii (Fig. 2), and, in contrast to domestic-ruminant density, which showed a negative relationship, these relationships were statistically significant. The statistically significant negative effect of season was linked to the higher risk of exposure to C. burnetii in the spring (Fig. 2). According to outputs from partial models (for unmanaged, managed, and farmed deer data sets) (Table 5), host and environmental factors were also evidenced as relevant drivers of exposure to C. burnetii. However, the main drivers for each particular management system differed. Whereas human density, season, and the red deer

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environmental favorability index were retained by the best-fitted risk factor model for

unmanaged deer, average spring temperature was retained by the best-fitted model for

managed deer, and season and domesticruminant density appeared to be the main drivers

of the risk of exposure to C. burnetii in red deer farms (Table 5; Fig. 2).

Figure 2. Relationships between the seroprevalence of C. burnetii in the population and explanatory factors identified through risk factor modeling for each of the modeled data sets (overall deer populations [OD], unmanaged populations [UD], managed populations [MD] and farmed populations [FD]).

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Discussion

This study constitutes a transnational-scale survey of C. burnetii in European wild ruminants and a first approach to identifying the factors that drive the ecology of C. burnetii in red deer. We found that C. burnetii is present in approximately 50% of the free-ranging and farmed Iberian red deer populations and that systemic infections occur in 50% of them. These facts support the involvement of the red deer in the ecology of C. burnetii. Indeed, to support the notion that a particular host species is acting as a true reservoir for a specific pathogen, provided that the species is well distributed and abundant (Wobeser, 1994), one must determine that (i) the pathogen is widely distributed in populations of that host within a relatively large territory, (ii) the pathogen is able to cause systemic infections (Maurin & Raoult, 1999; González-Barrio et al., 2015b), and

(iii) the host is able to shed the pathogen. We confirm the first two requisites here; the third requisite was confirmed previously (González-Barrio et al., 2015c). Thus, the red deer may be confirmed as a true C. burnetii reservoir.

Methodological considerations. The true seroprevalence of C. burnetii in free-ranging deer populations may have been underestimated because most deer sera (1,017 of 1,151) were collected from early autumn to midwinter, the main big-game-hunting season in

Iberia. Recent data from LO farm (Fig. 1) suggest that annual individual seroprevalence fluctuates according to the red deer calving season, with the lowest values in winter and maximumvalues in late spring (González-Barrio et al., 2014). Seroprevalence levels are higher in late spring to early summer, coinciding with the time of calving and, supposedly, with the main Coxiella burnetii excretion season.

Geographic distribution of C. burnetii in red deer populations.

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The wide geographical distribution of C. burnetii in Iberian red deer populations is noteworthy. To date, to the best of our knowledge, no exhaustive national study of C. burnetii in wild ruminants has been performed in Europe. C. burnetii DNA was found in the tissues of 23% of the roe deer analyzed from 9 of the 12 Dutch provinces during the massive human Q fever epidemic affecting The Netherlands from 2007 to 2010 (Rijks et al., 2011); however, the number of samples analyzed was low (n=79). Comparisons with results from previous regional studies of Spanish red deer are difficult because of differing geographic scales and techniques for diagnosing pathogen exposure: an indirect immunofluorescence test by which 9.5% of wild red deer and 40.0% of farmed red deer were found to be seropositive (Ruiz-Fons et al., 2008b) and molecular analyses (Astobiza et al., 2011a) where none of the 22 red deer analyzed tested positive. In general terms, on the premise of possible underestimation of real seroprevalence values, we may conclude that C. burnetii circulates widely in Iberian red deer populations.

Factors modulating the exposure of red deer to C. burnetii.

The modeling output of the overall deer data set partly confirmed our second hypothesis; environmental and host factors were found to be significant drivers of C. burnetii transmission in red deer. However, no effect of the management system on the risk of exposure to C. burnetii was observed, although the main drivers in partial data sets differed slightly (Table 5). The three management categories of red deer considered in this study are related to deer abundance and aggregation (Acevedo et al., 2008; Gortázar et al., 2006). Therefore, according to the observed effect of cattle density on the risk of exposure to C. burnetii (Álvarez et al., 2012; Piñero et al., 2014), we expected a clear effect of management. One would expect that in deer farms, and even in some intensively managed free-ranging deer populations, horizontal C. burnetii transmission would be enhanced due to the high animal-to-animal contact rate. This seems not to occur in general

119 terms, perhaps due to the complexity of C. burnetii ecology and the existence of multiple reservoir hosts (Ruiz-Fons, 2012; González-Barrio et al., 2015b). The low percentage of variance explained by the best-fitted model for unmanaged deer populations (Table 5) may reflect the existence of a complex scenario in environments with higher biological diversity. This would suggest that endemic cycles of C. burnetii implicating different wild

(and domestic) host species might be established in Iberia.

Climatic conditions during the C. burnetii shedding season modulate the risk of exposure of deer to this pathogen. Nonetheless, partial models revealed that the average spring temperature is relevant only in free-ranging managed populations; this variable itself explained 16.5% of model variance. Whether this observation is related to a direct effect of temperature on C. burnetii survival or transmission, or to indirect, nonconsidered effects— e.g., an effect on transmission— cannot be determined with our findings and with the existing literature. Therefore, this finding should be the basis for further studies aiming to deepen in C. burnetii ecology.

Although a general effect of the season was evidenced, the risk of exposure to C. burnetii was higher in the spring for unmanaged deer and similar in the spring and winter for farmed deer. This observation for farmed deer may be caused by a seasonal bias in farmed deer sampling in this study; only deer from the LO farm, which had a high proportion of seropositive deer, were surveyed in the winter. The observation for unmanaged deer agrees with the higher level of shedding of C. burnetii expected at the time of deer calving in midspring, as mentioned above.

Finally, host effects were revealed by the general model and by models for unmanaged and farmed deer populations. The influence of human density in the general model may be slightly modulated by an apparently exceptional result (Fig. 2) from a red deer

120 population with high seroprevalence. However, this factor also modulated the exposure of unmanaged deer to C. burnetii, thus showing the influence of human activities on the risk of exposure of deer to this pathogen. The density of coexisting domestic ruminants seems to dilute the risk of exposure of deer to C. burnetii. This result is shocking for a pathogen that is endemic in domestic ruminants in Iberia and whose transmission has been proven to be linked to host density (Álvarez et al., 2012; Piñero et al., 2014). This contrasting finding again suggests that the ecology of C. burnetii in wildlife is complex and that different wild and domestic species are involved in its maintenance, independently of the ability of the red deer to act as a true reservoir host. Indeed, the modeling output suggests that although an independent cycle of C. burnetii in red deer is posible without the intervention of a third species (susceptibility, systemic infection, and shedding demonstrated), other hosts may be implicated in the circulation of C. burnetii in wild foci.

Implications for animal and human health.

Haydon et al. (2002) redefined the reservoir concept for multihost pathogens and stated that a specific pathogen of relevance for a target host of interest may be maintained through a high number of combinations of host populations or environments that keep the pathogen circulating. Therefore, defining the risk of transmission of C. burnetii from red deer to target hosts (livestock and humans) is difficult, preventing us from concluding whether the red deer plays a major role in C. burnetii maintenance in Iberia or not. We believe that red deer populations constitute a highly relevant node in the life cycle of C. burnetii, but particular scenarios of interaction with third species need to be further investigated. Wild lagomorphs and small mammals infected by C. burnetii, among others, may excrete infectious bacteria (González-Barrio et al., 2015b; Barandika et al., 2007;

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Thompson et al., 2012) and therefore constitute relevant pieces of the C. burnetii maintenance and transmission puzzle.

The risk of C. burnetii transmission from red deer to humans could be comparable to that from livestock if deer-human and livestock-human indirect interaction rates were similar.

This is supported by the fact that both individual and population seroprevalences in red deer are similar to those found in domestic ruminants (Álvarez et al., 2012; Hamlin et al.,

2000; Astobiza et al., 2012a). Most effective livestock-human C. burnetii transmission events occur indirectly, through aerosols (Maurin & Raoult, 1999). We may expect that most deer-livestock and deer-human transmission events would occur indirectly

(Kukielka et al., 2013). Therefore, the risk of transmission from deer to livestock and humans depends on the exposure rate of deer, suggesting that extensively produced domestic ruminants and humans involved in hunting and wild ungulate management and conservation face a higher risk (Tozer et al., 2014; Whitney et al., 2009).

Our results point to free-ranging deer, perhaps in connection with other wild and domestic hosts, and deer farms as the main hot spots of circulation of C. burnetii among red deer in Iberia, and perhaps elsewhere in Europe. Further clarification of particular red deer- livestock or red deer-human interaction rates at different geographic scales should improve the chances of preventing C. burnetii transmission events.

Acknowledgments

We are grateful to game estate owners, gamekeepers, natural and national park managers, and farm managers for their collaboration in sample collection. Special thanks go to José

Antonio Ortiz from the LO farm and to Christian Gortázar for substantial support.Wealso acknowledge the great efforts of colleagues from the SaBio group at IREC in sample

122 collection (Joaquín Vicente, Pelayo Acevedo, Isabel G. Fernández-de-Mera, Ursula

Höfle, Paqui Talavera, Óscar Rodríguez, Álvaro Oleaga, Diego Villanúa, Vanesa Alzaga,

Elisa Pérez, Raquel Jaroso, Raquel Sobrino, Encarnación Delgado, Jesús Carrasco,

Ricardo Carrasco, Rafael Reyes García, Pablo Rodríguez, Mauricio Durán Martínez,

Valeria Gutiérrez, and many others). This work was funded by EU FP7 grant ANTIGONE

(278976) and CDTI (Centro para el Desarrollo Tecnológico Industrial, Spanish Ministry for the Economy and Competitiveness [MINECO]). J. A. Barasona holds an FPU predoctoral scholarship from MECD. J. P. V. Santos and J. Queirós were supported by

Ph.D. grants (SFRH/BD/65880/2009 and SFRH/BD/73732/2010, respectively) from the

Portuguese Science and Technology Foundation (FCT). F. Ruiz-Fons is supported by a

“Ramón y Cajal” contract from MINECO.

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Capítulo II. 2

124

Estado de Coxiella burnetii en las poblaciones de conejo de monte

(Oryctolagus cuniculus) en la península ibérica y factores de riesgo

asociados

European Rabbits as Reservoir for Coxiella burnetii

David González-Barrio, Elisa Maio, Madalena Vieira-Pinto, Francisco Ruiz-Fons

Emerging Infectious Diseases. 2015. 21 (6): 1055-1058

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Resumen

En este trabajo estudiamos el papel que juega el conejo de monte (Oryctolagus cuniculus) como reservorio de Coxiella burnetii en la región Ibérica. Altas seroprevalencias tanto individuales como en las poblaciones son observadas en conejos silvestres y granjas de conejos, la evidencia de infecciones sistémicas y la excreción vaginal apoyan el papel de reservorio de Coxiella burnetii al conejo de monte.

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Abstract

We studied the role of European rabbits (Oryctolagus cuniculus) as a reservoir for

Coxiella burnetii in the Iberian region. High individual and population seroprevalences observed in wild and farmed rabbits, evidence of systemic infections, and vaginal shedding support the reservoir role of the European rabbit for C. burnetii.

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Wildlife play a major role in the maintenance and transmission of multihost pathogens

(Ruiz-Fons et al., 2008c; Viana et al., 2014). Understanding the role of host species involved in multihost zoonotic pathogen maintenance and transmission is essential to prevent disease caused by these pathogens.

Coxiella burnetii, which is the cause of Q fever, is a zoonotic pathogen that infects multiple hosts (Maurin & Raoult, 1999). The implication of wildlife in the life cycle of

C. burnetii has been reported worldwide (Ejercito et al., 1993; Ruiz-Fons et al., 2008b), and wildlife might act as a source for humans infections (Marrie et al., 1986; González-

Barrio et al., 2015c).

European rabbits (Oryctolagus cuniculus) are native to the Iberian Peninsula and have been introduced into Australia, New Zealand, Chile, and Argentina (Monnerot et al.,

1994). Domestic varieties of European rabbits are farmed worldwide. Specific ecologic traits (high density, gregarious behavior, high reproductive rate) suggest that these rabbits might become a major reservoir of zoonotic pathogens. However, whether C. burnetii can infect, replicate in, and be shed by European rabbits and contaminate the environment is not known. In this study, we investigated the role of these rabbits in a region to which Q fever is endemic.

The Study

Serum samples were collected from European wild rabbits in 13 locations in Spain,

Portugal, and the Chafarinas Islands during 2003–2013 (Figure 1). Wild rabbits from 1 of the study locations (LO; Figure 1) were obtained from 2 epidemiologic scenarios (Maio et al., 2011). The first scenario involved rabbits that coexisted with farmed red deer

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(Cervus elaphus) (sites A and B). The second scenario involved rabbits that had not been in contact with ruminants since 2002 (site C).

In addition to serum samples, spleen, uterus, and mammary gland samples and vaginal and uterus swab specimens were collected from rabbits surveyed at location LO. Each rabbit from this location was weighed and sexed. Serum samples were also collected from farmed rabbits on 4 farms in Spain (Figure 1). Samples were stored at -20°C until tested.

Serum samples were analyzed by using the LSIVet Ruminant Q Fever Serum/Milk

ELISA Kit (Life Technologies, Carlsbad, CA, USA) and horseradish peroxidase– conjugated protein G (Sigma-Aldrich, St. Louis, MO, USA) as secondary antibody (Maio et al., 2011). Results were interpreted according to manufacturer’s recommendations.

DNA from tissues and swab specimens was extracted bu using the DNeasy Blood and

Tissue Kit (QIAGEN, Hilden, Germany). Swabs were incubated at 56°C for 30 min in

200 μL of AL buffer containing 20 μL of proteinase K. Swabs specimens were then vortexed for 15 s and removed. The remaining solution was incubated at 56°C for 30 min.

The manufacturer’s blood extraction protocol was then used. DNA aliquots were frozen at –20°C. Negative controls (nuclease-free water; Promega, Madison, WI, USA) were included during DNA extraction.

DNA samples were analyzed by using a conventional PCR (Berri et al., 2000). PCR products were visualized by electrophoresis in 1.2% agarose gels containing 0.1 μL/mL of GelRed Nucleic Acid Gel Stain (Biotium, Hayward, CA, USA).

Logistic regression models were used to test the effect of potential factors (Table) on the individual risk of exposure to C. burnetii. Individual ELISA results were included as response variables in the models and the location origin of rabbits was used as a random factor.

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Logistic regression models were also used for individual exposure of rabbits from location LO to C. burnetii (ELISA), for the presence/absence of C. burnetii DNA in spleen (a proxy of systemic infection), and for the presence/absence of C. burnetii DNA in the reproductive tract (a proxy of shedding; including PCR results from uterus, and vaginal and uterus swab specimens). Location LO was included as a random factor, and sex, weight and ruminant presence/absence were also included as predictor variables

(Table). Models were created by using a forward stepwise procedure. The model with the lowest Akaike information criterion (Akaike, 1974) was selected.

Statistical analyses were performed in SPSS version 20.0 (IBM, Armonk, NY, USA).

Prevalence-associated, Clopper-Pearson exact 95% CIs were estimated.

Serum samples from 572 rabbits (464 wild and 108 farmed) (Figure 1) were analyzed.

Overall seroprevalence rabbits, 37.9% (95% CI 33.5%–42.5%) for wild rabbits, and 8.3%

(95% CI 3.8%–15.2) for farmed rabbits. Seroprevalence in wild rabbit populations ranged from 6.7% to 81.3%. Nine (64.3%) of 13 wild rabbit populations and 2 (50%) of 4 farms had >1 seropositive rabbit. The best model for C. burnetii exposure retained sampling year and season, and the risk for seropositivity was higher in summer (Table).

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Figure 1. Seroprevalence of Coxiella burnetii (sample size) in wild and farmed European rabbits

(Oryctolagus cuniculus), Iberian Peninsula and Chafarinas Islands. The distribution area of wild rabbits in the Iberian Peninsula (10 x 10 km Universal Transverse Mercator squares) is shown

(gray shading) according to Mitchel-Jones et al. (1999). LO sampling location is indicated.

*Rabbit farm.

Seroprevalence at location LO was 65.2% (133/204; 95% CI 58.2%–71.7%); it was slightly lower at site C than at sites A and B (Figure 2, panel A). However, none of the considered factors were retained in the best model (Table). Six (4.4%; 95% CI 1.6%–

9.4%) of 136 spleen samples analyzed at location LO were positive by PCR (4 male and

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2 female rabbits). Five of the 6 spleen PCR–positive animals were seropositive. The 2

female rabbits were positive for C. burnetii DNA in vaginal swab specimens. Spleen

PCR–positive rabbits were observed only at sites A and B (Figure 2, panel B).

Table. Varibles considered as potential risk factors and outputs (coeficient/statistic) of best fitted risk factor models for Coxiella burnetii exposure in European rabbits (Oryctolagus cuniculus), Iberian Peninsula and Chafarinas Island*

Variable code Variable, units Cbsp CbspLO CbsplLO CbrtLO

Intercept NA 67.776/4.98† -037270.00‡ -2942.687/1.15‡ 2925.025/0.49‡ X Longitude, decimal degrees § ¶ ¶ ¶ Y Latitude, decimal degrees § ¶ § ¶ Ye Year -0.033/0.20‡ § 1.464/0.42‡ -1.453/0.45‡ Se Season § § § § Sp Spring 1.209/5.45‡ § -1.583/2.78‡ § Su Summer 2.257/5.45‡ § Referent § Au Autumn 0.043/5.45‡ § § § Wi Winter Referent § § § Mg Management, wild vs. farmed § ¶ ¶ ¶ Rum Ruminants, presence vs. absence ¶ § 0.059/0.0‡ 2.004/1.08‡ Sex Sex, M vs. F ¶ § -0.383/0.27‡ 2.004/0.22‡ Wg Weight, g ¶ § § §

* Cbsp, overall seropositivity; CbspLO, seropositivity at location LO; CbspILO, systemic infection (C.

1 burnetii DNA in spleen of wild rabbit); CbrtLO, shedding (C. burnetii DNA in reproductive tracts

of wild rabbits); NA, not applicable. †p≤0.05.

‡p˃0.05. §Variable was included in each model but was not retained in the best model.

¶Variable not tested.

The best model for the presence of C. burnetii DNA in spleen retained sampling year,

season, presence of ruminants, and sex (Table). Results suggest expected higher systemic

infection prevalence in rabbits coexisting with farmed red deer (Figure 2, panel B). C.

burnetii DNA was detected in the reproductive tract of 9 (14.1%; 95% CI 6.6%–25.0%)

of 64 female rabbits at sites A, B, and C (Figure 2, panel F). The presence of ruminants

was retained in the best model for C. burnetii DNA in the reproductive tract (Table). None

of the 13 mammary glands analyzed was positive for C. burnetii DNA.

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Conclusions

This study provides 3 results that suggest that European rabbits might be reservoirs of C. burnetii. These 3 results are high seroprevalence of this bacteria; systemic infections; and bacterial shedding in vaginal secretions, which, in other host species, constitutes the main source for environmental contamination and transmission between species (Guatteo et al.,

2007).

Figure 2. Prevalence of antibodies against Coxiella burnetii and C. burnetii DNA in European rabbits (Oryctolagus cuniculus) at sampling location LO, Iberian Peninsula. A) Antibodies; B) DNA in spleen; C) DNA in vaginal swab specimen; D) DNA in uterine swab specimen; E) DNA in uterus; F) DNA in reproductive tract (vaginal swab specimen, uterine swab specimen, uterus). Gray bars indicate seroprevalence. St_P indicates results for sites with ruminants (sites A and B); no ruminants were present at site C. Values at the top of bars indicate number of samples, and values at the bottom of bars indicate number of positive samples. Error bars indicate prevalence- associated exact 95% CIs.

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Host density is a major factor in C. burnetii prevalence in livestock (Piñero et al., 2014).

The highest seroprevalence values were observed at 2 locations where rabbit populations are managed for hunting purposes, which promotes high densities of rabbits. These findings suggest that rabbit density may be a major factor in the ecology of C. burnetii.

In addition, the European rabbit is a gregarious species with a high reproductive rate. This rate favors transmission of C. burnetii from infected to susceptible animals, which is enhanced by replacement of C. burnetii–negative rabbits and can contribute further to spread of this bacterium in the environment.

The higher risk of exposure to C. burnetii observed during the summer might be related to increased indirect interaction with C. burnetii shed by coexisting ruminants, whose main shedding season is late spring–early summer (Maurin & Raoult, 1999). Inclusion of ruminants in the final models for systemic infection and vaginal shedding at location LO might support this hypothesis. However, further analyses, including molecular typing of circulating strains, would be needed to confirm the direction, frequency, and magnitude of interspecies interactions favoring transmission of C. burnetii.

Indirect transmission of C. burnetii between rabbits, humans, livestock, and other wild species may be enhanced in regions with high-density rabbit populations and in regions in which the European rabbit is a major game or farm species. Hunters, game keepers, rabbit farmers, veterinarians, wildlife researchers, livestock producers and livestock might be exposed to C. burnetii from rabbits (Marrie et al., 1986; Whitney et al., 2009).

The European rabbit shows a high potential as a reservoir of C. burnetii for infection of livestock and humans in Europe.

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Acknowledgments

We thank location LO farm managers, Tania Carta, María Martín, Christian Gortázar and

José Antonio Ortiz for assistance during the study and Ursula Höfle for checking the

English grammar of the paper.

This study was supported by European Union FP7 grant ANTIGONE (278976), European

Union FP7 EMIDA ERA-NET grant APHAEA on wildlife disease surveillance in

Europe, and Centro para el Desarrollo Tecnológico Industrial, Spanish Ministry for

Economy and Competitiveness. F.-R-F. was supported by Juan de la Cierva and Ramón y Cajal contracts from the Spanish Ministry for Economy and Competitiveness.

Mr. González-Barrio is a doctoral student at the Spanish Wildlife Research Institute,

Ciudad Real, Spain. His research interests are the epidemiology of pathogens transmitted between wildlife, livestock, and humans within a OneHealth approach; the epidemiology and diagnosis of C. burnetii infections; and development of infection control strategies for wildlife.

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Capítulo II. 3

136

Dinámica de la infección por Coxiella burnetii en una población

endémica de ciervo en condiciones semi-extensivas

Long-term dynamics of Coxiella burnetii in farmed red deer (Cervus elaphus)

David González-Barrio, Isabel G. Fernández-de-Mera, José Antonio Ortiz, João Queirós, Francisco Ruiz-Fons

Frontiers in Veterinary Science. Veterinary Infectious Diseases. 2015. Aceptado

137

Resumen

Muchos aspectos de la dinámica de Coxiella burnetii que son relevantes para la implementación de estrategías de control en rebaños de rumiantes donde esta bacteria es endémica son desconocidos. Se diseñó un estudio en el tiempo para supervisar la dinámica de exposición de Coxiella burnetii en un rebaño endémico de ciervos a fin de permitir el diseño de métodos de control específicos de la fiebre Q. Otros aspectos relevantes en la dinámica de Coxiella burnetii como el efecto del estado inmune del rebaño; la edad, estación e infección temprana sobre la exposición; la vida media de anticuerpos frente a Coxiella burnetii; la presencia y duración de la inmunidad humoral maternal y la edad de la primera exposión fueron analizados. La dinámica de Coxiella burnetii en rebaños de ciervos parece etar modulada por factores del rebaño e individuales y en particular por las características de vida del hospedador. Las ciervas están expuestas a Coxiella burnetii al incio de su segundo año de vida ya que los anticuerpos maternales las protegen desde su nacimiento hasta después de la estación principal de excreción del patógeno que es a final de primavera principio de verano. La presión de infección varía entre años, probablemente asociada al efecto de inmunidad del rebaño, determinado variaciones inter-anuales en el riego de exposición. Estos resultados sugieren que cualquier estrategia aplicada para el control de Coxiella burnetii en rebaños de ciervos deden ser diseñados para inducer la inmunidad en su primer año de vida inmediatemente depués de la pérdida de los anticuerpos maternales. La vida media corta de los anticuerpos frente a Coxiella burnetii sugiere que cualquier protección inducida por la inmunidad humoral podría requerir una revacunación cada 6 meses.

138

Abstract

Several aspects of the dynamics of Coxiella burnetii that are relevant for the implementation of control strategies in ruminant herds with endemic Q-fever are unknown. We designed a longitudinal study to monitor the dynamics of exposure to C. burnetii in a red deer herd with endemic infection in order to allow the design of Q fever specific control approaches. Other relevant aspects of the dynamics of C. burnetii - the effect of herd immune status, age, season and early infection on exposure, the average half-life of antibodies, the presence and duration of maternal humoral immunity and the age of first exposure - were analysed. The dynamics of C. burnetii in deer herds seems to be modulated by host herd and host individual factors and by particular host life history traits. Red deer females become exposed to C. burnetii at the beginning of their second year since maternal antibodies protect them after birth and during the main pathogen shedding season - at the end of spring-early summer. Infection pressure varies between years, probably associated to herd immunity effects, determining inter-annual variation in the risk of exposure. These results suggest that any strategy applied to control C. burnetii in deer herds should be designed to induce immunity in their first year of life immediately after losing maternal antibodies. The short average life of C. burnetii antibodies suggests that any protection based upon humoral immunity would require revaccination every 6 months.

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Introduction

Coxiella burnetii is a worldwide-distributed gram-negative intracellular bacterium that causes Q fever, a zoonotic disease shared by humans and animals. Infection with C. burnetii in humans is usually asymptomatic but it may trigger acute and chronic clinical manifestations. Coxiella burnetii is also one of main pathogens causing reproductive losses in livestock (Oporto et al., 2006) and reproductive failure in pets (D'amato et al.,

2014; Kosatsky et al., 1984) and wildlife (Lloyd et al., 2010; Kersh et al., 2010;

Kreizinger et al., 2015; González-Barrio et al., 2015c). Clinical signs of Q fever in domestic ruminants are diverse; it has been associated with sporadic cases of abortion, premature delivery, stillbirth and weak offspring in cattle, sheep and goats, but epidemics with increased reproductive failure have been reported for sheep and goats mainly

(Agerholm, 2013). Since Coxiella burnetii infection does not always manifest clinically, the extent of C. burnetii infection in animals is probably underestimated.

Exposure to C. burnetii is increasingly reported in wildlife, e.g.: i) White-tailed deer -

Odocoileus virginianus - in the eastern US (Kirchgessner et al., 2013); ii) Rats - Rattus norvegicus and R. rattus - in the UK (Meredith et al., 2014) and the Netherlands (Reusken et al., 2011); or iii) European rabbits - Oryctolagus cuniculus - and Eurasian wild boar -

Sus scrofa - in the Iberian Peninsula (Astobiza et al., 2011a; González-Barrio et al.,

2015b; González-Barrio et al., 2015d). Recently, González-Barrio et al. (2015a) found that C. burnetii circulates endemically in Iberian red deer populations. This study suggests that the red deer plays an important role in the maintenance of C. burnetii in Europe. Thus the analysis of the dynamics of C. burnetii in red deer may be of interest to prevent Q fever transmission at the wildlife-livestock-human interface (Gortázar et al., 2014).

Furthermore, the presence of C. burnetii in red deer may have implications for red deer itself and coexisting wild species (Lloyd et al., 2010; González-Barrio et al., 2015c;

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Chaber et al., 2012; Hussein et al., 2012). Q fever may be an important cause of reproductive losses in red deer farming (González-Barrio et al., 2015c), an activity that is increasing worldwide (Hoffman & Wiklund, 2008). Therefore, deer producers could be interested in implementing Q fever prevention and control measures that would benefit from knowledge of the effect of deer farming particularities on C. burnetii dynamics.

Information on the dynamics of C. burnetii in endemic ruminant herds and on driving factors (host population, host individual and environmental) is scarce. A trade-off between infection pressure and herd immunity may influence infection dynamics in C. burnetii endemic herds, which may modulate the efficiency of vaccination trials. Recently it has been postulated that in endemic dairy cattle herds the immune status of the population drives exposure to C. burnetii (Piñero et al., 2014). According to this postulate, high levels of protection in an endemic herd may lead to a reduction in environmental contamination with C. burnetii, therefore reducing transmission. However, as long as the immune status of the herd changes with time (i.e. herd immunity decreases due to culling of immune individuals) while C. burnetii persists in latently infected animals or in infected fomites (Maurin & Raoult, 1999), the circulation of C. burnetii reactivates and expands within the population. Currently, no long time series study has demonstrated that the immune status of a C. burnetii endemic ruminant population changes with time to support this postulate. Information from long time series would provide a significant boost to understand the epidemiology of Q fever and plan any prevention and/or control approach.

Apart from host population factors, host individual factors (e.g. age, maternal-derived immunity o acquired immunity, among others) may modulate the dynamics of C. burnetii

(González-Barrio et al., 2015a). Currently, the presence, prevalence and duration of maternal anti-Coxiella burnetii antibodies and their effect in the outcome of natural

141 exposure to C. burnetii are poorly understood. If vaccination of animals at early ages

(before natural infection by C. burnetii takes place) is to be performed (Astobiza et al.,

2011b), knowledge on the exact timing for vaccination – i.e. the time at which maternal antibodies disappear and prior to exposure to C. burnetii - could be paramount to warrant protection. In dairy cattle, the transmission of colostral antibodies to calves borne from seropositive cows has been reported (Tutusaus et al., 2013) but the duration of these antibodies was not monitored. Could early exposure to C. burnetii of individuals in endemic herds modulate infection with the individual’s age? The effect of early exposure to C. burnetii on future protection against infection is also poorly known. In natural infections in domestic ruminant females, a non-immune animal is supposed to become infected and undergo a primary subclinical infection at early ages (Woldehiwet, 2004) that reactivates during the first pregnancy. Understanding the effect of natural early exposure to C. burnetii on future exposure would perhaps allow predicting the effect of vaccination at early ages on protection against C. burnetii infection. The likelihood of becoming infected by C. burnetii increases with age (Ruiz-Fons et al., 2010c). Indeed, age-related C. burnetii serological patterns have been reported in domestic ruminants

(Capuano et al., 2004; Guatteo et al., 2007) with highest seroprevalence in cows and sheep aged 3–5 years. Is this pattern similar in farmed red deer?

In this study we aimed to answer different questions that are relevant to understand C. burnetii dynamics in endemic ruminant herds in the long-time scale and that are, therefore, essential for the efficient planning and application of any Q fever control measure such as vaccination. The following objectives were addressed in the study: 1)

Analysis of long-time variation in exposure to C. burnetii; 2) Determination of the effect of herd immune status on the risk of exposure to C. burnetii of yearling females; 3) Test of the effect of deer age on exposure and humoral response to C. burnetii; 4) Study of

142 the effect of deer life-history traits (i.e. concentrated calving) on exposure to C. burnetii;

5) Investigation of the effect of natural exposure to C. burnetii at early ages over the future dynamics of exposure; 6) Determination of the average half-life of antibodies against C. burnetii; 7) Estimation of the presence, prevalence and duration of maternal antibodies; and 8) Determination of the age at which deer get exposed to C. burnetii for the first time in their life. To achieve our objectives a C. burnetii endemic red deer herd was selected as model. Therefore, the present study provides a case report on the dynamics of C. burnetii in a red deer farm with a history of C. burnetii infection in humans and reproductive failure in deer.

Materials and methods

Study farm and management schemes

The study was performed in a semi-extensive red deer farm located in the province of

Cádiz (Southern Spain) that was found consistently positive to C. burnetii in consecutive studies (González-Barrio et al., 2015c; Ruiz-Fons et al., 2008b). The number of deer on the farm was approximately 500 females and 80 males along the study period, with only slight inter-annual variations in the number of reared animals. The deer are semi- extensively bred within large (6-8 has) enclosures separated by high-wire fencing in batches of approximately 60-80 females; males are kept in separate enclosures. The habitat in the deer enclosures consists on patches of natural Mediterranean scrubland - mainly composed by evergreen (Quercus ilex) and cork (Quercus suber) oak trees - with large areas of year-round irrigated prairies. Animals are bred in separate but contiguous batches according to sex and breeding status. Artificial feed is provided daily on the farm.

The deer are managed from two to four times per year at maximum to avoid excessive stress. Management is carried out for sanitary issues, weaning, reposition and artificial

143 insemination. Hinds give birth naturally - i.e. without human intervention - in hidden areas of scrubland patches within enclosures from the end of April to the beginning of

May. Calves are weaned in August at 3.5 months of age and thereafter are kept in male and female batches in separate enclosures. Ear tagging is performed at weaning for identification of individuals. Calves are managed again at 7 months of age for sanitary reasons. When animals are 13 months old, a selection of yearling females and males for farm reposition is performed and the rest of yearlings are sold. Reposition yearling females are inseminated at the age of 16 months when they join other hinds in existing batches of reproductive females. Selected yearling males are kept separate from stags in reproductive condition. Adult deer, both males and females, are managed for sanitary control in January and in August each year. Reproductive females (>16 months old) are annually managed for artificial insemination in September.

Reproductive hinds in the study farm are culled annually according to their reproductive fitness or health status. Average productive life of deer females in the farm is unknown; some hinds remain productive for 13-15 years but most are culled at 4-5 years of life.

Management schemes in the farm are scheduled to carry out batch, reproductive and sanitary management issues without inducing excessive stress in the deer that, although farm-bred, still behave like wild animals. Therefore, animal sampling could only be performed according to the management schedule of the farm except for the monitoring of antibodies in the 2013 cohort (see details in the following section).

Survey design

We designed a retrospective survey to search for the presence and the level of antibodies against C. burnetii in deer sera collected in the study farm for disease surveillance purposes. The retrospective survey was aimed at testing for variation in herd

144 seroprevalence over time (Objective 1), the effect of herd immune status on the risk of exposure of yearling females (Objective 2), age-related variation in immune status of individual deer (Objective 3), seasonal variation in infection risk (Objective 4), influence of exposure at early ages on future exposure to C. burnetii (Objective 5) and the average half-life of C. burnetii antibodies (Objective 6). Blood samples were collected according to farm disease surveillance schemes and in the framework of Spanish and EU laws for notifiable disease surveillance. Therefore, no Animal Ethics Committee approval was required for the collection of blood samples from deer for the retrospective study.

To determine the inter-annual variation in exposure to C. burnetii we carried out a selection of deer sera (yearling - 12-24 months old - and adult - >24 months old - females; n=1021) collected in the farm along 12 consecutive years (2003-2014). Minimum annual sample size was estimated with WinEpi (http://winepi.net/sp/index.htm). The estimated sample size for each age-class - yearling and adult - with a 95% confidence level for a minimum seroprevalence of 5% with an accepted 10% error was 19; this minimum sample size was covered each year for each deer age-class. Selection of sera was carried out according to batch origin to obtain a balanced subset of samples that provides representative information of the real status of C. burnetii prevalence in the herd. For homogeneity of results, only sera collected in summer were selected to estimate inter- annual variation in the humoral immune status of deer in the farm.

To test for the rest of aforementioned objectives (2-6), we selected sera from reposition females belonging to three different cohorts - animals born in 2008, 2009 and 2010. Blood was collected from these females at least in four consecutive occasions (i.e. from 7 to 27 months old) and up to 13 consecutive occasions (i.e. from 7 to 78 months old). Deer were surveyed in summer and winter each year. Serum samples were obtained for the 2008 cohort at 7, 13, 20, 27, 32, 38, 44, 51, 56, 62, 67, 73 and 78 months of life. The same

145 survey was carried out for the 2009 cohort but up to 67 months of life and the 2010 cohort could be surveyed up to the 56th month of life. Minimum sampling size at each survey time was estimated with WinEpi employing the same parameters described above.

Although culling with age reduced the number of available samples with individuals’ age, sampling size was above 19 for each month class except for individuals at 73 and 78 months of age (n=13 in both cases).

Finally, to test for the presence, prevalence and duration of maternal antibodies (Objective

7) and for the age at which deer get exposed to C. burnetii for their first time in life

(Objective 8), serum samples from 21 calves born in 2013 were prospectively collected at 2, 3, 7, 13, 14, 19 and 20 months of life. This subset of the 2013 cohort was specifically surveyed to achieve these objectives since blood from calves in the farm is normally first collected at 7 months of age. The Research Ethics Commission of Castilla – La Mancha

University Animal Ethics Committee (Spain) approved this research.

Since we initiated an experimental vaccination trial in the study herd in January 2012 with a C. burnetii phase I inactivated vaccine, animals that were included in this study for

2012, 2013 and 2014 were exclusively unvaccinated animals of the control group. The interpretation of the evolution of the immune status of the herd will be carried out in this study considering this potential confounding factor for 2012-2014 results.

Serological analyses

Serology has been widely employed to test for the status of C. burnetii in ruminant populations (Ruiz-Fons et al., 2010c; Ruiz-Fons et al., 2008b) and to understand its dynamics even though a proportion of infected individuals may not seroconvert (De

Cremoux et al., 2012). In a vaccination trial in the study deer farm (the authors, unpublished) near the 90% of vaccinated seronegative deer seroconverted after a single

146 dose of a C. burnetii inactivated phase I vaccine. This percentage was close to the 100% after a boosting dose 3 weeks later. Therefore, in contrast to domestic ruminants (e.g. sheep, Astobiza et al., 2011b), it is expected that a high percentage of infected red deer display detectable levels of anti-C. burnetii antibodies in ELISA. Therefore, ELISA was employed to study the dynamics of infection by C. burnetii.

Blood was collected from the jugular vein into sterile tubes and it was thereafter kept at

4º C and transported to the laboratory. Blood was centrifuged at 3,000g for 10 minutes and the serum obtained was preserved at -20ºC until analyses were performed. The presence of specific antibodies against C. burnetii phase I and II antigens in deer sera was determined with a commercial indirect ELISA test (LSIVet™ Ruminant Q Fever

Serum/Milk ELISA Kit, Life Technologies, USA) with an in-house modification in the secondary antibody (Protein G−Horseradish peroxidase, Sigma-Aldrich, USA) that had been validated for wild ungulates (González-Barrio et al., 2015a). ELISA results were expressed as the sample-to-positive control ratio (SP). For each sample, the SP was calculated according to the formula:

"SP ="(("ODs - ODnc" ))/(("ODpc - ODnc" ))"x100" where ‘ODs’ is the optical density of the sample at a dual wavelength 450-620 nm,

‘ODnc’ is the optical density of the negative control and ‘ODpc’ is the optical density of the positive control. All SP values ≤40 were considered as negative, whereas SP values

>40 were considered as positive. The SP ratio was considered as a proxy of the level of antibodies against C. burnetii as suggested by the manufacturer.

Statistical analyses

Statistical analyses were carried out to test different hypotheses: i) the immune status of the herd is negatively related to the incidence of infection by C. burnetii in yearling

147 females (Objective 2); ii) the age of individuals is positively related to seroprevalence and antibody levels (Objective 3); iii) there are seasonal variations in the rate of exposure to

C. burnetii (Objective 4); and iv) variations in the immune status at early ages modulates future exposure to C. burnetii (Objective 5).

To assess for the effect of herd immune status on the incidence of C. burnetii in yearling females, we performed Spearman correlations with the annual incidence

(presence/absence of C. burnetii antibodies) in yearlings as response variable and three different explanatory variables that were tested separately: i) seroprevalence in adult females in the same year (t); ii) seroprevalence in adult females in the previous year (t-

1); and iii) average seroprevalence in adult females in years t, t-1 and t-2 (two years before). Seroprevalence in adult females was employed as a proxy of herd seroprevalence because adult females constitute around the 75% of the herd. Spearman correlations were also employed to test the relationship between antibody levels and individuals’ age.

Mann-Whitney U non-parametric tests for independent samples were employed to test for the alternative hypothesis of statistically significant differences in average antibody levels by season. Chi-square tests were employed for the same purpose with seroprevalence as response variable.

Finally, the influence of the immune status of individuals at early ages (7 months old; presence/absence of C. burnetii antibodies) on the evolution of the humoral immune response against C. burnetii infection along individual’s age was tested by repeated measures ANOVA. Individual squared SP values were transformed into natural logarithms for normality and were employed as response variable. For this analysis, we used data obtained from the 2008, 2009 and 2010 deer cohorts.

148

IBM SPPS v22.0 (IBM, Armonk, NY, USA) was employed for statistical analyses. Exact

Clopper-Pearson 95% confidence intervals (95%CI) were estimated for prevalence values

using Quantitative Parasitology 3.0 software (http://www.zoologia.hu/qp/qp.html).

Results

A total of 373 (inter-annual sampling size range: 19-92) and 648 (inter-annual sampling

size range: 19-159) serum samples collected in summer from yearling and adult females,

respectively, were selected for the cross-sectional approach during 2003-2014. The inter-

annual evolution of herd (yearling+adult) and age class-specific seroprevalence from

2003 to 2014 is shown in Figure 1.

Figure 1. Evolution of herd (yearling+adult) average and age class-specific (yearling - 12-24 months old - and adult - >24 months old) seroprevalence (and associated 95% exact confidence intervals) from 2003 to 2014.

Average annual herd and age-specific seroprevalence values varied between years. There

was no clear time scale pattern in inter-annual herd seroprevalence that remained above

30% and below 70% along the study period. Seroprevalence in adult females fluctuated

149 between years with periods of 1 to 3 consecutive years of high values (above 70%) followed by periods of 1 to 2 consecutive years of medium values (from 30% to 60%).

The high seroprevalence in unvaccinated females in 2012, 2013 and 2014, even though

>70% of the herd had been vaccinated (the authors, unpublished), indicate a natural period of high herd humoral immunity similar to that observed in 2005-2007. A unimodal pattern was observed in yearlings. The observed age-related pattern of humoral immunity in individual deer shows that we can assimilate annual seroprevalence in yearlings to annual incidence ratio. Incidence in yearlings was low (5-10%) from 2005 to 2008 (although samples were not analysed in 2006), it increased (35-43%) from 2008 to 2012 and it steeply decreased (25% in 2013 and 6% in 2014) from 2012 onwards.

Incidence in yearlings remained low during the periods in which high seroprevalence was observed in adult females and increased during a period in which the seroprevalence in adult females was lower than in preceding years. Incidence was nonetheless not statistically influenced by seroprevalence in adult females at times t (survey year) and t-

1 (previous year to survey), and by average seroprevalence in adult females in years t to t-2. However, trends in all three relationships were negative, that is increasing incidence in yearlings related to decreasing seroprevalence in adults in the same or in previous years.

One thousand four hundred and forty-five sera from 217 animals born in 2008 (n=97),

2009 (n=92) and 2010 (n=28) were sequentially analysed from 7 (calf) up to 78 (adult) months of life to estimate the evolution of antibodies against C. burnetii with age. Not every animal could be surveyed sequentially along the study period because of annual culling for health or productive reasons. On average, both seroprevalence and the level of antibodies in individual deer increased with age and this pattern was evidenced in animals from the three cohorts (Figure 2). Age (in months) and the level of antibodies were

150 statistically significantly correlated (rho=0.416, p<0.001). This result confirms increasing levels of antibodies with deer age, but highest antibody levels were observed between 4 and 5 years of life.

Figure 2. Age-related evolution of C. burnetii level of antibodies (A) and seroprevalence (B) for

2008, 2009 and 2010 deer cohorts together. Season is depicted in the x-axis of each chart (W: Winter; S: Summer). Exact 95% confidence intervals for seroprevalence and standard error for average antibody levels are displayed in the charts.

The monitoring of individual deer sera by ELISA showed an evident seasonal pattern in antibody levels and seroprevalence. Both the level of antibodies and seroprevalence peaked in summer each year and decreased through winter (Figure 2). Seasonal

151

differences were statistically significant: i) Average SP in summer was 73.3±2.5 in

contrast to 37.8±1.4 in winter (Z=-12.89, df=1, p<0.001); and ii) 67.3% (95%CI: 62.8-

71.7) of animals surveyed in summer had antibodies in contrast to 36.0% (95%CI: 32.6-

39.5) in winter (X2=110.93, df=1, p<0.001). This pattern was evident for any of the three

cohorts surveyed (Figure 2).

Results from the repeated measures ANOVA showed that differences in the evolution of

the level of antibodies in relation to the presence/absence of C. burnetii antibodies at 7

months of age were not statistically significant (Figure 3). Around the 82% of individuals

in the 2008-2010 cohorts were seronegative at 7 months of age.

Figure 3. Evolution of antibody levels (and associated standard error) with individuals’ age according to the presence/absence of anti-C. burnetii antibodies by ELISA (S/P>40) at 7 months of age for 2008, 2009 and 2010 deer cohorts together.

152

The evolution of the presence and level of antibodies in calves born in 2013 from two to

20 months of life are shown in Figure 4. High levels of antibodies were evidenced in calves at 2 months (June 2013), when 75% of them displayed an SP ratio >40 (i.e. seropositive). Thereafter, both the level of antibodies and seroprevalence decreased sharply in one month (July 2013) and disappeared at 7 months (November 2013). Animals remained seronegative at 13 months of age (May 2014) and then became seropositive at

14 months (June 2014), two months after the calving season in the farm. This seroconversion was most probably caused by natural infection and affected 50% of the animals. The average level of antibodies derived from natural infection at 14 months was lower than that acquired from their mothers during lactation (at 2 months); in seropositive animals average SP ratio was 122.4 at 2 months (n=15) in comparison to 77.8 in seropositive animals at 14 months of life (n=7). Both seroprevalence and antibody level remained at similar values at 19 months (November 2014) but decreased thereafter notably a month later (December 2014). This observation and the seasonal pattern observed indicate that the expected average life of antibodies against C. burnetii could be around 6 months. These results show that deer become exposed to C. burnetii for the first time in life mainly at around 12-14 months of age.

Figure 4. Age-related evolution of C. burnetii level of specific antibodies (A) and seroprevalence (B) in farmed red deer (2013 cohort). Exact 95% confidence intervals for seroprevalence and standard error for average antibody levels are displayed in the charts.

153

Discussion

Understanding the factors that drive the dynamics of endemic pathogens is paramount to design and efficiently apply any preventive or control measure. Vaccination with phase I inactivated C. burnetii vaccines - one of the main Q fever control tools in domestic ruminants (Van de Brom et al., 2015) - is recommended for naïve or low-prevalence herds

(Guatteo et al., 2008) but not for endemic herds (Astobiza et al., 2011b; Astobiza et al.,

2011c). However, the status of C. burnetii in an endemic host population may present inter-annual variation (Piñero et al., 2014) with years in which the percentage of naïve individuals in the population is high, followed by years in which this percentage is low.

Identification of time windows with high percentage of naïve individuals in a C. burnetii endemic population would allow the implementation of vaccination trials. During these time windows the percentage of susceptible individuals that could be protected by vaccination would be enhanced. Determining the factors that modulate the dynamics of

C. burnetii in endemic populations would allow predicting the occurrence of appropriate windows to implement vaccination. This study improves our understanding of the dynamics of C. burnetii in endemic ruminant herds and driving factors that could allow more efficient control approaches in the future.

Risks of infection by C. burnetii associated to red deer

Red deer may be a relevant reservoir of C. burnetii in Europe because of its increasing relevance as a game resource, its current demographic status and the status of C. burnetii in red deer populations. Deer farming is increasing (Hoffman & Wiklund, 2008) and likewise populations of free-roaming red deer currently display increasing demographic trends (Apollonio et al., 2010). Additionally, free-roaming deer populations are increasingly managed as extensively bred ruminants (supplementary feeding, fencing,

154 translocations) but lacking appropriate sanitary control (Gortazar et al., 2006). Changes in livestock production schemes in the Netherlands - increasing number of goat herds without C. burnetii control - led to the 2007-2010 epidemics of human Q fever (Roest et al., 2011a), demonstrating how important demographic changes of a single host may be to increase the risk of infection by C. burnetii. Interestingly, farmed and free-roaming

Iberian red deer populations display similar population and individual seroprevalence values to livestock (González-Barrio et al., 2015a; Woldehiwet, 2004). Furthermore, increasing geographic distribution and population density of red deer in Europe may increase the implication of this wild species in future Q fever epidemics. Prevention would only be possible if accurate scientific knowledge is available. Our study provides insights into poorly studied epidemiological aspects of the dynamics of C. burnetii in red deer populations.

Long-term dynamics of C. burnetii in farmed red deer

There is a main question to answer in relation to the dynamics of C. burnetii: Does the status of C. burnetii change with time in an endemic ruminant herd? Our results for 12 consecutive years show fluctuation in the status of C. burnetii prevalence in a ruminant herd in which the pathogen circulates endemically. Piñero et al. (2014) also provided evidence of inter-annual variation in endemic dairy cattle herds but within a shorter time period.

Inter-annual variation of the status of an endemic pathogen in a herd could be the consequence of the trade-off between pathogen burden and host immunity if we assume that there are no changes in the composition of the herd - e.g. size, culling and import rates, age and sex structures - and the influence of external pathogen sources remains constant (González-Barrio et al., 2015d; Piñero et al., 2014). Those features remained

155 constant along the study period in the deer farm. Even the potential influence of other sources of C. burnetii such as European rabbits - Oryctolagus cuniculus - remained similar along the study period, with C. burnetii seroprevalence from 2005 to 2013 above

50% (González-Barrio et al., 2015b). Therefore, herd immunity effects seem to be the most probable cause of changes in the epidemiological status of C. burnetii in the herd.

However, in spite of the observed negative trend in the relationship between incidence in yearling females - probably associated to infection pressure - and seroprevalence in adult females, relationships were not statistically significant. Increasing incidence in yearling females from 2009 to 2012 coincided with a period of lower seroprevalence values in adult females in comparison to 2005-2007. In contrast, there was a steep decrease in incidence in yearlings from 2012 to 2014 when seroprevalence in adult (non-vaccinated) females was again high. This observation could potentially be linked to herd vaccination against C. burnetii (the authors, unpublished data, Capítulo IV) or alternatively be the consequence of a natural period of high humoral immunity in the herd, but in any case it indicates variation in infection pressure. Interestingly, an outstanding rate of reproductive failure in the herd in 2011 - that was presumably caused by Q fever (González-Barrio et al., 2015c) - coincided with the increasing incidence observed in yearlings in 2009 to

2012.

It was unfortunately impossible to monitor the presence and burden of C. burnetii in the environment in the farm (aerosols, soil, water, food, pastures) along the study period in order to accurately estimate the evolution of infection pressure with time. However, this has been carried out in dairy cattle and changes in the epidemiological status of C. burnetii in dairy cattle herds are linked to the detection of C. burnetii in manure, air and dust samples (Piñero et al., 2014). According to Piñero et al. (2014) high herd humoral immunity levels would reduce shedding of large burdens of C. burnetii to the

156 environment, therefore reducing infection pressure. This would result in a reduced incidence in naïve (yearling) individuals in the herd. Another observation that may support variation in infection pressure with time is that 30.9% (30/97) and 9.9% (9/91) of deer calves born in 2008 and 2009, respectively, were seropositive to C. burnetii at 7 months of age (naturally infected in their first year of life) in contrast to 0% (0/21) in

2013. This pattern paralleled observations in incidence in yearling females. These changes could be linked to the implementation of vaccination in the herd from 2012 onwards since vaccinated and unvaccinated deer coexist in existing enclosures in the farm.

Short-term herd effects on the dynamics of C. burnetii in red deer

Intra-annual variation in exposure to C. burnetii has been previously suggested in wildlife studies (Pioz et al., 2008b). Ruminant females shed C. burnetii mainly around parturition and therefore in species with a defined breeding season shedding should be concentrated.

In contrast to dairy cattle, which breed along the year, the breeding season of the red deer is concentrated at the end of spring (Clutton-Brock et al., 1982). This fact implies that, within a year, there is a predominant shedding season during which the risk of exposure of individuals is higher. The short half-life of C. burnetii antibodies allowed differentiating that the risk of exposure in winter is much lower than by the end of spring- early summer, which is consistent with a predominant shedding season in deer than coincides with the breeding season. This particularity of the epidemiology of C. burnetii in farmed red deer may favour the implementation of control strategies since adequate management measures in liaison with medical treatments can significantly reduce the exposure of individuals around the breeding season.

Host individual traits influencing C. burnetii dynamics in red deer

157

Host individual traits may modulate the relationship that a host establishes with C. burnetii and age-related effects have been described frequently (Ruiz-Fons et al., 2010c).

In this study we found a significant increase in seroprevalence and antibody level with the individuals’ age. This may be caused by cumulative effects of continuous exposure to

C. burnetii with time or may be linked to increasing immune competence with the individual’s age. Two observations point to an effect of host immune competence as the causal factor for this age-related increase in seroprevalence and antibody levels: i) the increasing trend in both parameters up to the 4th year of life (similar to findings in cattle,

Guatteo et al., 2008) and the decreasing pattern thereafter; and ii) the low average half- life of anti-C. burnetii antibodies observed (discussed below).

Acquired immunity after natural infection by C. burnetii at early ages may have a protective effect over the outcome of future infections since reproductive failure caused by Q fever is more evident in primiparous females and decreases with age (Astobiza et al., 2011b; Astobiza et al., 2011c). However, we observed that in calves exposed to C. burnetii at 7 months of age the average humoral immune response induced by infections in adulthood did not differ from that observed in non-exposed calves at that age. Whether the effect derived from natural infection is similar to what we would expect from vaccination is difficult to predict, but this finding suggests that acquired immunity at early ages does not prevent re-infection by C. burnetii in the future. This could be linked to the short average half-life of C. burnetii antibodies observed in deer. Vaccination of deer at early ages with an appropriate re-vaccination calendar would perhaps induce long-lasting protection against infection by C. burnetii.

The pattern of antibody levels in the 2013 cohort suggest that deer calves get antibodies from their mothers early in their lives that then disappear before their 7th month of life.

Maternal-acquired antibodies have also been reported from cattle calves (Tutusaus et al.,

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2013). Dairy cows infected with C. burnetii maintain detectable levels of antibodies along the gestation period and even after partum (García-Ispierto et al., 2011) that are transmitted to new-born calves with the colostrum. Whether maternal antibodies protect against infection by C. burnetii is unknown. Results from the 2013 cohort suggest that maternal antibodies protect calves in their first year of life - perhaps in association to the concentrated shedding season in late spring, but the presence of antibodies in 7 month- old animals of the 2008/2009 cohorts contradicts that observation. If we assume - on the basis of incidence rates - that infection pressure was higher in 2008/2009 than in 2013 and that a high percentage of calves born every year acquire maternal antibodies, we may hypothesize that under high infection pressure in the herd a percentage of the calves are not protected during their first year of life. Only proper experimental approaches with a controlled challenge would offer information to understand the effect of humoral immunity on protection (Roest et al., 2013a). Nonetheless, our findings suggest that deer calves should be vaccinated for the first time when they are around 5-6 months of life.

The exact timing for vaccination should be determined through future experiments with a higher sampling frequency.

An interesting finding that should be born in mind when planning vaccination protocols in deer farms is that any protection linked to humoral immunity would last only around 6 months. Maternal antibodies in the 2013 cohort were high at 2 months of age and completely disappeared 5 months later. This observation and the sharp decrease of antibodies from natural infection from months 14-19 to month 20 suggest an average half- life of anti-C. burnetii antibodies of 5-to-6 months without natural re-infections.

Therefore, re-vaccination every 6 months would be recommendable to maintain humoral immunity in deer. The average half-life of antibodies in other species may be higher since

159 antibodies can be detected even a year after infection in humans (Angelakis & Raoult,

2010).

Conclusions

Red deer are able to maintain C. burnetii and transmit it to other wildlife, livestock, pets and humans. Current knowledge on the status of C. burnetii in red deer in Iberia together with results obtained in this study point to this species as a source of Q fever that needs to be considered by animal and public health authorities.

In endemic herds C. burnetii inter-annual dynamics may be modulated by host herd and individual factors that should be considered for planning efficient control approaches.

Particular host life history traits (e.g. concentrated breeding) also have an important effect on the intra-annual variation in the dynamics of C. burnetii. Naturally acquired humoral immunity seems to have no effect on future re-infection of deer by C. burnetii, perhaps linked to the observed short average half-life of antibodies in red deer.

Acknowledgements

We are grateful to people that contributed to sample collection, including farm keepers and members of the SaBio group at IREC (Tania Carta, Joaquín Vicente, Mauricio

Durán and Óscar Rodríguez). We thank Dr. Ursula Höfle for reviewing the English grammar and expression of the manuscript. The research was funded by ‘Centro para el

Desarrollo Tecnológico Industrial’ (CDTI) of the Spanish Ministry for the Economy and Competitiveness (MINECO) and by EU-FP7 ‘Antigone’ project. FRF acknowledges funding by the ‘Ramón y Cajal’ programme of MINECO. IGFM is funded by the University of Castilla – La Mancha.

160

Capítulo II. 4.

161

Genotipos de Coxiella burnetii presentes en fauna silvestre en la

península ibérica basados en MLVA

Coxiella burnetii genotypes in Iberian wildlife

David González-Barrio, Ferry Hagen, Jeroen JHC Tilburg, Francisco Ruiz-Fons

FEMSE Microbiology Ecology. En revisión.

162

Resumen

Para investigar si los genotipos de Coxiella burnetii, agente causante de la fiebre Q, que circulan en la fauna silvestre están asociados con los genotipos que infectan a ganado doméstico y a humanos, se llevó a cabo el análisis mediante MLVA (Muliple-locus variable number tándem-repeat de muestras procedentes de ciervo (Cervus elaphus), jabalí (Sus scrofa), conejo de monte (Oryctolagus cuniculus), rata de campo (Rattus norvegicus) y ratón de campo (Apodemus sylvaticus). El tipado mediante MLVA fue realizado usando 6 loci variables en Coxiella burnetii: Ms23, Ms24, Ms27, Ms28, Ms 33 y Ms 34. La base de datos de Coxiella burnetii dentro de MLVABank 5.0 fue utilizada para comparar los genotipos encontrados en este estudio con los 344 genotipos de divero origen dentro de esta base de datos. 22 genotipos de fauna silvestre y dos genotipos de cabra doméstica fueron identificados. Algunos genotipos identificados en fauna silvestre fueron también encontrados en en casos de fiebre Q en humanos, sugeriendo que los humanos y la fauna silvestre comparten genotipos de Coxiella burnetii. El genotipado muestra una baja diversidad en genotipos de Coxiella burnetii dentro de la misma especie de hospedador que entre hospedadores. Estos resultados proveen importantes conocimientos para enternder la epidemiologia de Coxiella burnetii en la interfaz fauna- ganado-humano.

163

Abstract

To investigate if Coxiella burnetii, the causative agent of Q fever, genotypes circulating in wildlife are associated with those infecting livestock and humans, multiple-locus variable number tandem-repeat (MLVA-6-marker) analysis was carried out over isolates from red deer (Cervus elaphus), Eurasian wild boar (Sus scrofa), European wild rabbit

(Oryctolagus cuniculus), brown rat (Rattus norvegicus) and wood mouse (Apodemus sylvaticus). MLVA typing was performed by using six variable loci in C. burnetii: Ms23,

Ms24, Ms27, Ms28, Ms33 and Ms34. The C. burnetii cooperative database from

MLVABank 5.0 was employed to compare genotypes found in this study with 344 isolates of diverse origin. Twenty-two genotypes from wildlife and two genotypes from domestic goats were identified. Some MLVA genotypes identified in wildlife were also isolated from human Q fever clinical cases, suggesting that humans and wildlife share C. burnetii genotypes. Genotyping showed lower within-host than between-host diversity of C. burnetii genotypes. These results provide important insights to understand the epidemiology of C. burnetii at the wildlife-livestock-human interface.

164

Introduction

Coxiella burnetii is the causative agent of Q fever, a zoonosis that affects humans and mammals worldwide (Angelakis & Raoult, 2010). During the past decade information on the epidemiology, pathogenicity and control of Q fever in European domestic ruminants, which constitute the major C. burnetii reservoirs, has grown in the scientific literature

(Angelakis & Raoult, 2010). In parallel, genotyping studies have been performed mainly as a cause of the large-scale human Q fever outbreak in the Netherlands from 2007 to

2010 (Roest et al., 2011a; Tilburg et al., 2012b). In the European context, C. burnetii genotypes that circulate among domestic ruminants and humans in Hungary, Poland,

Portugal, Spain and the Netherlands have been genotyped (Astobiza et al., 2012b;

Chmielewski et al., 2009; Jado et al., 2012; Santos et al., 2012; Roest et al., 2013b; Sulyok et al., 2014; Tilburg et al., 2012a). In contrast, information regarding the relevance of wild hosts in the ecology of C. burnetii is scarce (González-Barrio et al., 2015a;

González-Barrio et al., 2015b; González-Barrio et al., 2015c; González-Barrio et al.,

2015d) and, consequently, genotypes circulating in wildlife have been rarely identified

(Jado et al., 2012, Rijks et al., 2011). Interestingly the origin of several human Q fever cases remains unclarified (EFSA 2014a) and human-wildlife interaction has been suggested as a risk factor for human infection with C. burnetii (Whitney et al., 2009). As long as the efficiency and the range of application of C. burnetii control measures in domestic animals increase (Astobiza et al., 2011c; Piñero et al., 2014), wild reservoirs of

Q fever may become more relevant (EFSA 2010). Therefore, from an epidemiologic perspective, typing wildlife-associated C. burnetii genotypes will contribute to identify the epidemiological link, if any, between wildlife and human and/or livestock Q fever cases. This information would improve currently on-going prevention and control measures (e.g. by improving biosafety tools and protocols).

165

Genotyping C. burnetii from wildlife will help tracing back clinical cases in humans directly exposed to wildlife, e.g. hunters, wildlife keepers, veterinarians and slaughterhouse staff, as well as of people exposed indirectly (e.g. by wildlife-generated infected aerosols). Therefore, the aim of the current study was to type C. burnetii genotypes circulating in Iberian wildlife by applying multiple-locus variable number tandem-repeat analysis (MLVA). Typing allowed comparing molecular patterns of wildlife C. burnetii with patterns of genotypes infecting humans and domestic animals.

The information provided in this study is of great relevance to understand the epidemiology of Q fever.

Table 1. Details of host origin, geographic origin, type of sample and collection year of qPCR

positive samples considered for Coxiella burnetii genotyping in this study.

166

Materials and methods

Samples

Two hundred and fifteen C. burnetii real-time PCR (qPCR) positive samples obtained from spleen, milk or swabs of various wild species (red deer Cervus elaphus, Eurasian wild boar Sus scrofa, European wild rabbit Oryctolagus cuniculus, brown rat Rattus rattus and wood mouse Apodemus sylvaticus) and three goat (Capra aegagrus hircus) bulk-tank milk samples were included in this study. Details on the number of samples per geographic origin, collection year, sample type and host are shown in Table 1. Spleen samples were collected from free-ranging red deer harvested by hunters in commercial hunting events. Deer samples from Cádiz province - milk and vaginal swabs - were collected after 3-4 months from calving at a deer farm with a previous history of Q fever clinical cases in humans and suspected reproductive failure in deer caused by infection with C. burnetii (González-Barrio et al., 2015c). All samples from free-ranging wild

European rabbits and free-ranging wild boar were collected from animals harvested by hunters. Small mammals were captured with Sherman traps (H.B. Sherman Traps Inc.,

Tallahassee, FL, USA) and samples were collected during necropsies. European wild rabbits and small mammals from Cádiz province (Table 1) were collected in the surroundings or within the surveyed deer farm (González-Barrio et al., 2015b). Bulk-tank milk samples were collected from a goat farm located in the province of Sevilla, southern

Spain. The geographic locations of all samples are shown in Figure 1.

167

Figure 1. Map showing the geographic location of Coxiella burnetii qPCR positive samples

included in this study in relation to hosts. The number of samples per host and province is shown within each host drawing.

DNA extraction and qPCR

DNA from spleen, milk and swabs was extracted by using a commercial kit (DNeasy

Blood & Tissue kit, Qiagen, Hilden, Germany) following the protocols provided by the manufacturer

(http://mvz.berkeley.edu/egl/inserts/DNeasy_Blood_&_Tissue_Handbook.pdf). In order to improve DNA extraction from swabs, these were kept at 56ºC for 30 min in a solution containing 20 µl of proteinase K and 200 µl of AL buffer, vortexed for 15 sec and removed. The sample remained for 30 additional minutes at 56ºC; then, the

168 manufacturer's blood extraction protocol was followed. Each sample of milk (200 μl) was mixed directly with ATL and proteinase K and incubated for 3 h at 56ºC; then, the manufacturer's blood extraction protocol was followed. DNA aliquots obtained were quantified (NanoDrop 2000, Thermo Scientific, Waltham, MA, U.S.A.) and frozen at -

20ºC until PCR performance. In order to prevent and detect sample cross-contamination, negative controls (Nuclease free water; Promega, Madison, WI, U.S.A.) were included every 10 samples during the DNA extraction procedure. All samples were tested by a qPCR targeting the IS1111a insertion element of C. burnetii as described previously

(Tilburg et al., 2010).

Multiple-locus variable number tandem-repeat (MLVA) analysis

MLVA analysis was performed using 6 of the most variable loci of the 17 loci previous described (Arricau-Bouvery et al., 2006). We performed two multicolor multiplex PCR assays targeting six microsatellite markers containing either six or seven base pairs (bp) repeat units; 3 hexanucleotide repeat markers (Ms27, Ms28 and Ms34) and 3 heptanucleotide repeat markers (Ms23, Ms24 and Ms33). Primer sequences and PCR conditions have previously been described (Tilburg et al., 2012b, Klassen et al., 2009).

PCR was performed in a total volume of 20 µl containing 1 U of FastStart Taq DNA polymerase (Roche diagnostics, Almere, the Netherlands), 0.2 mM, dNTP´s, 4 mM

MgCl2 in 1× reaction buffer, 0.1-1.0 µM of amplification primers and 5 µl of DNA sample. Samples were 50 times diluted by using ddH2O. One µl of the diluted sample was added to 8.9 µl of water and 0.1 µl of CC-500 ROX internal size marker (Promega,

Madison, WI, USA). Analysis of the amplification products was performed on an

ABI3500xL Genetic Analyzer (Applied Biosystems, Foster City, CA, USA). DNA from the Nine Mile strain of C. burnetii (RSA 493) was used as reference. The number of repeats in each marker was determined by extrapolation using the relative size of the

169 obtained fragments to those obtained using DNA from the Nine Mile strain (9-27-4-6-9-

5 for markers Ms23-Ms24-Ms27-Ms28-Ms33-Ms34, respectively).

Table 2. Coxiella burnetii MLVA genotypes from Iberian wildlife and domestic goats obtained in this study.a MLVA type designation was given according ti host source (G: Goat; D: red deer and R: wild rabbit) and a numerical order. b The numbers of repeats in each MLVA locus was determined by correlating amplicon sizes with those obtained from Nine Mile strain (RSA493). Ø Almost complete MLVA profile.

MLVABank 5.0 as a tool to compare genotypes

The aim of MLVABank (http://mlva.u-psud.fr/mlvav4/genotyping/index.php) is to facilitate microbe genotyping (including pathogenic bacteria), essentially for epidemiological purposes. Data from a variety of assays can be managed, including polymorphic tandem repeat typing (MLVA), multiple locus sequence typing (MLST), single nucleotide polymorphisms (SNPs), and spoligotyping assays based upon clustered regularly interspersed palindromic repeats (CRISPRs). In our case, we used MLVABank

170 to compare our MLVA genotypes with previously published C. burnetii genotypes of diverse host and geographic origin. We used the Coxiella burnetii2014 cooperative database, which was set-up by the Centre National de la Recherche Scientifique (CNRS,

Paris, France) in 2014 by aggregating: i) MLVA data from published whole genome sequence data by in silico analysis; ii) Data published in 2006 by Arricau-Bouvery et al.

("C. burnetii 2007 Orsay" database); iii) Data, provided by Kinga Sulyok, Miklós

Gyuranecz and colleagues, Institute for Veterinary Medical Research, Budapest, Hungary

("C. burnetii 2014" Hungary” database); iv) Data, both published and unpublished, produced since 2007 by Jeroen Tilburg and co-workers ("C. burnetii 2014 Nijmegen" database); and v) Additional published data compiled from the literature ("C. burnetii published others" database).

Results

All 218 qPCR positive samples had a cycle threshold (Ct) value <40.0. Based on acquired experience only DNA-samples that revealed a Ct-value <35.0 were considered for MLVA genotyping. Twenty-six of these samples (9 from red deer, 3 from goats and 14 from rabbits) were considered for MLVA analysis (Table 2). Overall, 17 complete MLVA genotypes were obtained from 19 of the 26 samples tested. In 7 of the samples genotypes were almost complete, i.e. one of the 6 markers did not amplify (Table 2). Figure 2 shows the relationship between all identified genotypes from deer, rabbits and goats in this study.

Clustering of the MLVA genotypes using the minimum spanning tree method showed high diversity. In total, three different clusters were defined: i) Genotypes in cluster one were all obtained from deer and they are all interconnected by repeated number changes in one of the six markers; ii) Genotypes in cluster two were obtained from rabbits; and iii) Genotypes in cluster three included goat genotypes, which differ in four markers with

171 genotypes in cluster two. Clusters one and two are interconnected by three genotypes from rabbits that differ by repeated number changes in three of the six markers.

Interestingly, the wild rabbit genotype that differs in three markers with genotypes in the deer cluster comes from northern Spain (Table 2) whereas the rest of genotypes from rabbits come from the same location from which red deer genotypes were obtained.

Table 3. Samples analyzed for the comparative study in MLVABank based on the host (A) and geographic origin (B).

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Using MLVABank 5.0, we compared 370 genotypes, including the current samples

(Table 3). This comparison is based in the host origin (Appendix 1a) and in the geographic origin (Appendix 1b) of genotypes. The major part of rabbit genotypes clustered separately from red deer genotypes, except three rabbit genotypes. Ten of the rabbit

MLVA genotypes clustered with C. burnetii genotypes isolated from human cases in

Canada, France, Portugal and Spain. This rabbit-cluster is linked by a difference of only a single marker to human genotypes from Portugal (see Appendix 1b). The remaining rabbit genotypes (n=3) clustered together with the red deer genotypes in a cluster that included: i) C. burnetii isolated from ticks in the USA, including Nile Mile strains from

USA, Germany and the Netherlands; ii) humans from France, Poland, Portugal and Spain; iii) a Portuguese goat; iv) a Japanese dairy cow; and v) a hay sample from Sweden.

Discussion

Genotyping by MLVA was employed in this study because it has previously been proved to be a powerful method to type C. burnetii from a diversity of hosts and geographic origins (e.g. Roest et al., 2011b; Tilburg 2013). Therefore, MLVA is nowadays the first choice method to compare C. burnetii genotypes in spite of its limitations.

Few studies reported to have successfully obtained complete MLVA genotypes from wildlife; e.g. a study in the Netherlands could not obtain complete MLVA genotypes from roe deer (Capreolus capreolus; Rijks et al., 2011). Other authors opted for multiple PCR and hybridization methods to genotype C. burnetii of wild origin (Jado et al., 2012), making therefore comparison with MLVA results difficult. An additional constraint to typing C. burnetii in wildlife is the difficulty of surveying wildlife during the patent period, when replication of C. burnetii is higher. None of the wild animals surveyed in this study had symptoms of being infected by C. burnetii, which hinders obtaining samples with the adequate C. burnetii concentration for MLVA typing. The major part of

173 qPCR positive samples (181 out of 218) had Ct-values close to the negative threshold.

None of the MLVA genotypes isolated from rabbits in the current study has previously been described in other studies (Astobiza et al., 2012; Chmielewski et al., 2009; Santos et al., 2012; Sulyok et al., 2014; Tilburg et al., 2012b) perhaps indicating that a unique cluster of closely related genotypes is kept circulating in wild rabbit populations. In contrast, three MLVA genotypes isolated from red deer had been previously described in ticks, cattle and humans, other two genotypes (D14 and D15) have a similar MLVA profile to the genotype D1, which itself is identical to C. burnetii Nine Mile reference strain. MLVA genotypes D14 and D15 information of a single MLVA marker is missing

(for Ms33 and Ms34 respectively), resulting in an incomplete MLVA profile. If missing marker regions Ms33 (D14) and Ms34 (D15) contain nine and five repeat units respectively, their profile is identical to the C. burnetii Nine Mile reference strain. Most of the other deer genotypes differed only in one marker from those previously described.

This may perhaps suggest that these genotypes are microvariants of a founder genotype as suggested previously for genotypes from livestock (Astobiza et al., 2012b) or may be the result of misclassification by MLVA. Nonetheless, these results should be interpreted with caution due to the low number of C. burnetii that could successfully be genotyped in this study and the limited geographic diversity of origin of these genotypes. This drawback was perhaps caused by the low concentration of C. burnetii DNA in wildlife samples and the requirements of the typing method employed.

MLVA typing of C. burnetii from Iberian wildlife showed a lower diversity within-host species than between-host species, even though most of the genotypes came from red deer and rabbits closely coexisting within a C. burnetii endemic focus in southern Spain for which the presence of the pathogen has been demonstrated in consecutive surveys

(González-Barrio et al., 2015a; González-Barrio et al., 2015b; Ruiz-Fons et al., 2008b).

174

The three rabbit genotypes that clustered with deer genotypes (Fig. 2, Appendix 1b) suggest that inter-species transmission is feasible. However, the low number of similar genotypes found in coexisting deer and rabbits (Table 3, Fig. 2, Appendix 1b) suggests that inter-species transmission is not as frequent as it would be expected from the multihost nature of C. burnetii as previously suggested in coexisting domestic ruminant species

(see de Bruin et al., 2012). Interestingly, Iberian C. burnetii genotypes from rabbit and deer clustered mainly with C. burnetii isolated from human cases in France and Portugal, suggesting that a link between humans and wildlife in C. burnetii transmission exists as previously suspected (González-Barrio et al., 2015c; Whitney et al., 2009; Davoust et al.,

2014). This finding, in addition to the wide geographic distribution of exposure to C. burnetii of Iberian red deer and wild rabbit populations (González-Barrio et al., 2015b;

González-Barrio et al., 2015c) suggests that wildlife, red deer and European wild rabbit in particular, may be important wild sources of C. burnetii for humans. An additional interesting finding was the low similarity of MLVA genotypes from livestock and wildlife, even considering those from close geographic locations described in this study

(Table3); only genotypes from a Japanese cow and a Portuguese goat clustered with red deer and rabbit genotypes. This result could be perhaps associated to the design of the study since the coexistence of wildlife and livestock was not considered as a criterion for the survey, or could be certainly related to within host predominant circulation of C. burnetii genotypes as observed in coexisting deer and rabbits in this study and also in livestock (de Bruin et al., 2012). Further studies in wildlife-livestock interaction scenarios should be performed to assess whether coexisting wildlife and livestock share C. burnetii genotypes.

175

Figure 2. Minimum spanning tree showing the relationship between the obtained MLVA genotypes identified in this study and four sequenced Coxiella burnetii strains, i.e. Nine Mile RSA493

(AE016828), RSA331 (CP000890), CbuG Q212 (CP001020) and CbuK_Q154 were determined in silico using the published sequences (Tilburg et al., 2012a). Each circle represents a unique genotype; the size of the circle corresponds to the number of samples with that genotype. Complete

(19) and almost complete (7) MLVA genotypes were included in this analysis. Branch labels and connecting lines refer to the number of different markers between genotypes. Genotypes connected by a grey background differ in only one marker from each other and may represent microvariants of one founder genotype. Letters and numbers within the circles indicate species and genotype (“G” for goat, “R” for European rabbit and “D” for deer).

176

In conclusion, we obtained 15 complete and 7 almost complete C. burnetii genotypes from Iberian wildlife and two complete genotypes from domestic animals in spite of the abovementioned difficulties. Some of the MLVA genotypes identified are shared with humans, showing that the transmission of C. burnetii at the wildlife-human interface is possible. However, our results do not allow concluding any direction in C. burnetii transmission - from wildlife to humans or in the opposite direction. Red deer and

European rabbits should be considered as potential C. burnetii reservoirs in Iberia and, perhaps, also in the rest of Europe.

Funding

This work was funded by EU FP7 Grant ANTIGONE (278976) and CDTI (Centro para el Desarrollo Tecnológico Industrial, Spanish Ministry for Economy and

Competitiveness-MINECO). F.R-F is supported by the Ramón y Cajal program of the

Spanish Ministry for the Economy and Competitiveness and D.G-B acknowledges funding by Cátedra UCLM-Fundación ENRESA.

Acknowledgments

We are grateful to Rocío Jiménez Granado for providing the goat bull-tank milk samples and to colleagues at IREC for wild ungulate sampling. We also wish to thank farm keepers and the veterinarian - José Antonio Ortiz - of the Cádiz deer farm for their collaboration in sample collection.

177

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179

Capítulo II. 5.

180

Genotipado de Coxiella burnetii de fauna silvestre ibérica mediante

PCR e hibridación RLB y relaciones con genotipos de ganado

doméstico y humanos en España

Coxiella burnetii genotypes in Spanish wildlife: implications for livestock and

human health

David González-Barrio, Isabel Jado, Isabel G. Fernández-de-Mera, María Rocío

Fernández-Santos, Manuela Rodríguez-Vargas, Cristina García-Amil, Beatriz Beltrán-

Beck, Pedro Anda, Francisco Ruiz-Fons

181

Resumen

Las evidencias actuales apuntan a un papel relevante de la fauna silvestre en la ecología de Coxiella burnetii en todo el mundo. Sin embargo, la falta de información sobre los genotipos de C. burnetii circulantes en la fauna silvestre impide la trazabilidad de los casos de animales y de los casos clínicos de fiebre Q en humanos con un possible origen en la fauna silvestre. Por lo tanto, con el objetivo de comparar los genotipos de C. burnetii que circulan en España en la fauna silvestre, el ganado y los seres humanos, se genotiparon 87 muestras provenientes de fauna silvestre y 20 de Ganado mediante PCR-

RLB, de las cuales 38 eran de ciervo rojo (Cervus elaphus), 27 de conejo de monte

(Oryctolagus cuniculus), 13 de mapaches (Procyon lotor) y 9 de pequelos mamíferos

(Microtus arvalis y Mus spp. así como otras 2 muestras de cabra doméstica (Capra aegragus hircus) y 18 de oveja (Ovis aries). 90 de las 107 muestras fueron positivas a C. burnetii mediente PCR a tiempo real. Cuatro grupos genomics (I, II, VI y VII) se encontraron en fauna silvestre y otros cuatro en ganado (I, II, III y IV). Identificamos 7 diferentes genotipos, todos previamente descritos en España. Los genotipos encontrados en el ganado coinciden con los ya descritos dentro del grupo de ruminates domésticos. El genotipado mediante PCR-RLB confirma evidencias previas mediante genotipado por

MLVA, que sugieren que C. burnetii puede mostrar adaptaciones a especies particulares de hospedadores, ya que la mayoría de genotipos de conejo y ciervo se agrupan pore specie y separados entre sí. Los genotipos de fauna silvestre se agrupan principalmente con genotipos de garrapatas y con genotipos aislados de casos clínicos agudos de hepatitis en humanos. Estos resultados sugieren que determinados genotipos de C. burnetii pueden circular de forma natural en un ciclo entre garrapatas y fauna silvestre, y que ocasionalmente podrían afectar a humanos por la picadura de garrapatas o por exposición a la fauna silvestre. Este hallazgo puede estar detrás de la variación observada en la

182 presentación clínica de la fiebre Q aguda en humanos en España, con neumonía atípica predominante en el norte y en el sur de la hepatitis.

183

Abstract

Current evidences point to a relevant role of wildlife in the ecology of Coxiella burnetii worldwide. However, the lack of information on C. burnetii genotypes circulating in wildlife prevents tracing-back clinical animal and human Q fever cases with potential wildlife origin. Therefore, with the aim of comparing C. burnetii genotypes circulating in wildlife, livestock and humans in Spain, 87 samples from red deer (Cervus elaphus, n=38), European wild rabbit (Oryctolagus cuniculus, n=27), raccon (Procyon lotor, n=13) and small mammals (Microtus arvalis and Mus spp., n=9) as well as 20 additional samples from goat (Capra aegragus hircus, n=2) and sheep (Ovis aries, n=18) were genotyped by

PCR-RLB. Ninety of the 107 samples were positive to C. burnetii by qPCR. Four genomic groups - I, II, VI and VII - were found in wildlife and four - I, II, III and IV - in domestic ruminants. We identified 7 different genotypes, all previously described in

Spain. Livestock genotypes clustered mainly with previously reported genotypes in livestock. PCR-RLB genotyping confirmed previous findings from MLVA that suggest that C. burnetii may display adaptations to particular host species because most genotypes of sympatric deer and rabbits clustered in separate groups. Wildlife genotypes clustered mainly with tick genotypes and with genotypes isolated from acute hepatitis cases in humans. Those results suggest that particular C. burnetii genotypes may circulate naturally in a wildlife-tick cycle that may occasionally jump into humans through tick bites or exposure to wildlife. This finding may be behind the observed variation in the clinical presentation of acute Q fever in humans in Spain, with predominant atypical pneumonia in the north and hepatitis in the south.

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Introduction

Q fever is a zoonotic worldwide-distributed infectious disease caused by Coxiella burnetii, a gram negative and highly ubiquitous bacterium with high environmental resistance (Maurin & Raoult, 1999). Domestic ruminants (cattle, goat and sheep), pets

(cats and dogs) and, to a lesser degree, wild mammals are natural reservoirs of C. burnetii

(Angelakis & Raoult, 2010; González-Barrio et al., 2015a; González-Barrio et al.,

2015b). Transmission to humans occurs mainly at the livestock-human interface through aerosols contaminated with C. burnetii shed by domestic ruminats (Angelakis & Raoult,

2010). Coxiella burnetii infection of mammals is mostly assymptomatic, but in a low-to- moderate percentage of infections, acute and chronic courses occur. In humans, Q fever is associated with a multiple clinical spectrum, from asymptomatic or mildly symptomatic seroconversion to fatal disease. In acute cases, patients may present with any of the following clinical signs: fever, fatigue, chills, headache, myalgia, skin rash, sweats, nausea, vomiting, diarrhoea, cough, chest pain, pneumonia, hepatitis, myocarditis, pericarditis, meningoencephalitis and, even, death. A low percentage of acute cases - especially patients with previous valvulopathy and, to a lesser extent, immunocompromised persons and pregnant women - evolve to more severe and complicated chronic courses that may present with endocarditis, vascular alterations, osteoarticular disease, chronic hepatitis, chronic pulmonary infections or chronic fatigue syndrome which can be fatal without an appropriate treatment (Maurin & Raoult, 1999).

It is assumed that domestic ruminants are the main reservoir of C. burnetii for humans.

Identical genotypes have been isolated in humans, goats and sheep but not in cattle during the 2007-2010 Dutch outbreak of Q fever (Tilburg et al., 2012b). This finding has also been observed in Spain, where no genomic group (GG) of cattle has been found in humans

(Jado et al., 2012).

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Little information exists on the pathogenesis of C. burnetii strains associated with clinical cases in humans and animals (Van Schaik et al., 2013). Nevertheless, various studies have tried to identify molecular markers associated to the different clinical manifestations of acute and chronic Q fever in humans, which are also of application for the characterization of animal strains (Massung et al., 2012; Frangoulidis et al., 2013). Variations in plasmid

DNA of C. burnetii has been described to relate to clinical manifestations of infection

(Samuel et al., 1985). Plasmids QpH1 and QpRS were reported to be associated with acute and chronic Q fever, respectively, whereas plasmids QpDG and QpDV are not related with clinical manifestations. This plasmid classification is correlated with the genomic groups I to VI described after restriction fragment length polymorphism (RFLP) and microarray analyses (To el., 1998; Beare et al., 2006). QpH1 is present in genomic groups I-III. QpRS represents groups IV and V that have been isolated from patients with endocarditis (Maurin & Raoult, 1999). The group VI is formed by a special group obtained from feral rodents in Dugway (Utah, USA), which carries plasmids QpDG and

QpDV (Mallavia, 1991). Another molecular marker related to clinical manifestations of

C. Burnetii infection is the acute disease antigen A (adaA). This antigen was described as a diagnostic marker for acute Q fever (To et al., 1998; Zhang et al., 2005) whereas adaA negative strains are related to chronic cases.

In Spain, clinical manifestations are geographically dependent (Montejo et al., 1985), with a higher proportion of pneumonic forms in the north of Spain (Tellez et al., 1988;

Espejo et al., 2014) whereas in southern Spain the proportion of hepatitis and fever of intermediate duration cases is higher (de Alarcón et al., 2003). The reasons for this geographic clustering are still unclear and molecular studies would help clarifying this phenomenon. Jado et al. (2012) studied strains from human, livestock and wildlife origin and found that cattle seems not to participate in the transmission of C. burnetii to humans.

186

In contrast, sheep, goats, wild boar, rats and ticks share genotypes with the human population. None of the strains detected in wildlife (wild boar, rats and ticks) was among the most virulent genomics groups for humans (I-III) but clustered with strains isolated from human cases of acute hepatitis and chronic infections. The participation of wildlife in the life cycle of C. burnetii is a fact (Ruiz-Fons, 2012; González-Barrio et al., 2015a;

González-Barrio et al., 2015b) and wildlife-associated strains could carry genetic variations leading to potentially emerging strains causing trouble to animal and human health in the future (Gortázar et al., 2014a). Therefore, the aims of this study were: i) to characterize Coxiella burnetii genotypes circulating in Iberian wildlife; and ii) to estimate if wildlife genotypes are shared with domestic animals and humans in Spain.

Figure 1. Map showing the geographic location of samples included in this study for Coxiella burnetii genotyping in relation to hosts. The number of samples per host and province is shown within each host drawing

187

Material and Methods

Samples

One hundred and seven samples obtained from authoctonous Iberian wildlife, domestic animals and exotic wild species were collected for this study. Spleen samples of red deer

(Cervus elaphus) were collected from free-ranging red deer harvested by hunters in commercial hunting events in south-central (Ciudad Real province) and nothern Spain

(Asturias province). Deer samples from Cádiz province (southern Spain) - milk and vaginal swabs - were collected at a C. burnetii endemic red deer farm with a previous history of acute Q fever in humans and in deer (González-Barrio et al., 2015c). All samples from free-ranging European wild rabbits (Oryctolagus cuniculus) were collected from animals harvested by hunters in Cádiz, Sevilla (southern Spain), Ciudad Real,

Toledo (south-central Spain) and Zaragoza (north-eastern Spain) provinces. Samples from small mammals from Asturias and Palencia provinces (northern Spain) were collected during necropsies performed to individuals captured during previous projects of our group. Rabbits surveyed in Cádiz province were collected nearby the surveyed deer farm in the same province (González-Barrio et al., 2015a) where both species coexist.

Spleen samples from racoons (Procyon lotor) were collected during necropsies performed to individuals trapped in Madrid and Guadalajara provinces (central Spain) during population control campaigns approved by Spanish animal conservation authorities because of their invasive nature (Beltrán-Beck et al., 2012b). Samples collected from domestic animals consisted in bulk-tank milk samples from a dairy goat farm located in the province of Sevilla and semen samples from rams of the Manchega sheep breed in

Ciudad Real province (Ruiz-Fons et al., 2014b). Details on the number of samples per geographic origin, sample type and host are shown in Table 1. A map displaying the geographic locations of samples is shown in Figure 1.

188

DNA extraction

DNA from spleen, milk, swabs and semen was extracted by using a commercial kit

(DNeasy Blood & Tissue kit, Qiagen, Germany) following the protocols provided by the manufacturer

(http://mvz.berkeley.edu/egl/inserts/DNeasy_Blood_&_Tissue_Handbook.pdf). In order to improve DNA extraction from swabs, these were kept at 56ºC for 30’ in a solution containing 20 µl of proteinase K and 200 µl of AL buffer, vortexed for 15” and removed.

The sample remained for 30 additional minutes at 56ºC; then, the manufacturer's blood extraction protocol was followed. Each sample of milk (200 μl) was mixed directly with

ATL and proteinase K and incubated for 3 h at 56ºC; then, the manufacturer's blood extraction protocol was followed. DNA aliquots obtained were quantified (NanoDrop

2000, Thermo Scientific, Waltham, MA, U.S.A.) and around 200 ng of DNA were used in each PCR. To improve DNA extraction from semen we employed a protocol described previously (Ruiz-Fons et al., 2014b). In order to prevent and detect sample cross- contamination, negative controls (Nuclease free water; Promega, Madison, WI, U.S.A.) were included every 10 samples during the DNA extraction procedure. DNA extracted from COXEVAC vaccine (CEVA Santè Animale, France) was employed as positive control.

Molecular detection and genotyping of C. burnetii

Previous to genotyping a screening assay was used for the detection of C. burnetii;

IS1111-based PCR coupled with hybridization with a specific probe by reverse line blotting (RLB) was used for detection of C. burnetii (Willems et al., 1994; Berri et al.,

2000; Barandika et al., 2007; Jado et al., 2012). The reaction mix included 80 μg/tube of bovine serum albumin (Roche Diagnostics GmbH, Manheim, Germany), 3.75 mM MgCl2

(Applied Biosystems, Branchburg, NJ, USA) 200 μM dNTPs (Applied Biosystems,

189

Branchburg, NJ, USA) and 4U of AmpliTaq GoldW DNA Polymerase (Applied

Biosystems, Branchburg, NJ, USA). Primer concentrations ranged from 0.6 to 1 μM (Jado et al., 2012). The amplification cycles included an initial cycle of 94ºC for 9’, followed by 40 cycles of 94ºC 30”, 60ºC 1’, and 72ºC 1’, with a final extension at 72ºC for 10’.

The amplifications were performed in an MJ Research PTC-200 (Bio-Rad Laboratories,

S.A., Alcobendas, Spain) in volumes of 50 μl. Hybridization by RLB was performed as previously described (Jado et al., 2006) using 48ºC for the hybridization and 40ºC for the conjugate and the washing steps. Concentration of probes ranged from 0.8 to 6.4 pmol/μl

(Jado et al., 2012). Two overlapping films (SuperRX, Fujifilm España S.A., Barcelona,

Spain) were used in each assay to obtain a less and more exposed image for each membrane. The results of the GT study were further analyzed by using InfoQuest™FP

4.50 (BioRad, Hercules, CA, USA). Clustering analyses used the binary coefficient

(Jaccard) and UPGMA (Unweigthed Pair Group Method Using Arithmetic Averages) to infer the phylogenetic relationships.

Results

Ninety samples were positive to qPCR and RLB hybridization with cycle threshold (Ct) values <40.0. Six genomics groups were determined: I, II, III, IV, VI and VII as previously described (Jado et al., 2012). Four genomics groups - I, II, VI and VII - were found in wildlife samples, three - I, VI and VII - in red deer, two - I and VII - in wild rabbits and two - I and II - in racoons. In domestic animals four genomics groups - I, II,

III and IV - were found, three - I, II and III - in sheep and one - VI - in goat (Table 2).

Within genomic groups of red deer, group I was detected in three milk samples from

Cádiz province and in two spleen samples from Ciudad Real province. In other two samples of red deer from Cádiz - vaginal swabs and milk - genomics groups VI and VII

190 were detected. In rabbits, a total of 8 samples were genotyped. Group VII was detected in four samples from Cádiz (uterus and vaginal swabs) and in one sample (spleen) from

Toledo province. Two spleen samples of wild rabbits from Zaragoza and one spleen sample from Sevilla belonged to group I. Within the racoon samples (spleen), group I was detected in three samples from Madrid and group II in two samples from Guadalajara.

In the other hand, four different genomic groups were detected in livestock from Sevilla and Ciudad Real provinces. Groups I, II and III were found in semen samples of sheep in

Ciudad Real province. In goat bulk-tank milk samples from Sevilla province genomic group IV was found in two samples.

Finally, the adaA was present in all samples of genomic group III (all from sheep). Eigth of thirteen samples from group I were adaA positive. Samples of genomic groups II, IV,

VI and VII were all adaA negative (Table 2).

Table 1. Host origin, geographic origin, sample type and number of C. burnetii qPCR positive samples employed in this study.

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Table 2. Summary of genotypes within genomic groups found in domestic and wild animals analyzed in this study

*: Acute disease antigen A (adaA).

Host Sample Province Ct CBU0007 CBU0071 CBU0168 CBU0598 CBU0881 CBU1805 CBU2026 CBU0952* Genomic group

Goat Sevilla 31,12 IV- Bulk-tank milk (Capra aegagrus hircus) Sevilla 32,48 IV-

Guadalajara 34,54 I-

Madrid 35,08 I+ Racoon Spleen Madrid 35,27 (Procyon lotor) I+ Guadalajara 35,19 II- Guadalajara 35,46 II-

Cadiz 32,18 I+ Milk Cadiz 34,17 I+

Vaginal swab Cádiz 32,50 I+ Red deer Ciudad Real 33,02 I+ (Cervus elaphus) Spleen Ciudad Real 33,02 I+ Vaginal swab Cádiz 31,40 VI-

Milk Cádiz 33,22 VII-

Ciudad Real 35,29 I-

Ciudad Real 33,63 I+

Sheep Ciudad Real 34,23 II- Semen (Ovis aries) Ciudad Real 34,46 II-

Ciudad Real 35,29 III+

Ciudad Real 36,53 III+

Sevilla 35,76 I- Spleen Zaragoza 21,89 I- Zaragoza 25,12 I-

European wild rabbit Cádiz 28,48 VII- Vaginal swab (Oryctolagus cuniculus) Cádiz 31.18 VII-

Cádiz 29.93 VII- Uterus Cádiz 35,46 VII- Spleen Toledo 32,09 VII-

192

Discusion

In this study, we identify genotypes of C. burnetii present in wildlife by PCR and RLB hybridization and compare these with previously described genotypes in wildlife, humans and livestock in Spain. Information gathered in this study will be of help to understand the complex ecology of this multi-host pathogen in a country in which C. burnetii is endemic.

Methodological considerations

We opted for multiple PCR and hybridization previously described methods (Beare et al.,

2006; Jado et al., 2012) to genotype wildlife C. burnetii because of the higher amount of existing information in humans and livestock in Spain using this typing methods (Jado et al., 2012). Coxiella burnetii strains from wildlife have been previously genotyped by

MLVA (Rijks et al., 2011; Davoust et al., 2014; Cumbassá et al., 2015; Capítulo III.4).

However, MLVA information on Spanish C. burnetii genotypes is scarce at large scales in the country and multiplex PCR coupled with RLB hybridization is able to counteract some of the MLVA drawbacks such as the differentiation of microvariants of a same genotype (Piñero et al., 2015; Astobiza et al., 2012b; Capítulo III.4).

The method employed here is quick, reproducible and sensitive. It can be applied directly to clinical and environmental samples, and is able to identify up to 16 genotypes depending on adaA presence/absence. This will facilitate the acquisition of global data on the circulation of genotypes of C. burnetii and complements existing information obtained by MLVA. This method provides additional information on the virulence of C. burnetii strains of wildlife origin for humans since different genomic groups are involved in acute and chronic manifestations of Q fever. Van Schailk et al. (2013) proposed that genotypes in groups I, II and III are highly virulent and cause acute Q fever cases - the

Nine Mile strain is within group I – whereas the rest of genomic groups appear with higher

193 frequency in chronic Q fever cases. However, Jado et al. (2012) found most human acute

Q fever cases in Spain caused by genotypes VII and VIII and chronic cases associated to group IV. Other authors have described an outer membrane protein-coding gene (adaA) associated with acute Q fever-causing strains when present and to chronic cases when absent (Zhang et al., 2005). Few studies focused in wildlife genotypes of C. burnetii in

Spain with this method and therefore this study comes to complement existing information on wildlife.

Completing the spectrum of information on Coxiella burnetii genotypes in Spain

Overall, we identified 6 genomics groups and 7 different genotypes. All genotypes found in this study fell within previously described genomic groups (Hendrix et al., 1991;

Glazunova et al., 2005; Svraka et al., 2006; Arricau-Bouvery et al., 2006). All found genotypes except two were previously described in Spain (Jado et al., 2012). Genotypes found in goats and sheep in this study were previoulsy described in these species in Spain except genotypes I- and II- that were found in Manchega sheep from Ciudad Real province.

Genomic group I was found in sheep, racoons, deer and wild rabbits in this study.

Genotypes in this gemonic group have been isolated in humans with acute pneumonia and in a sheep placenta in northern Spain (Jado et al., 2012). Current results in Spain have found genotypes in group I in southern, central and northern Spanish provinces, suggesting this genomic group displays both geographic and host wide ranges in the country.

Genomic groups II and III have been detected in livestock previously (Beare et al., 2006;

Jado et al., 2012). However, although group III was herein detected only in sheep, genotypes in group II (specifically genotype II-) were detected in racoons. Whether racoons, an alochthonous and invasive species to Spain (Rodríguez-Refojos &

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Zuberogoitia, 2011), got infected from livestock or carried out this genotype from their areas of origin before introduction is difficult to determine. Previous reports in Spain found a restricted geographic distribution of group II genotypes; just genotype II+ was found in sheep in northern Spain (Jado et al., 2012). We detected the presence of genotype

II- in sheep and racoons in south-central Spain but not in the rest of studied provinces, suggesting a restricted distribution of C. burnetii genotypes within the genomic group II.

In contrast, Jado et al. (2012) described genotype III+ in samples from domestic ruminants allover Spain, including the province in which this genotype was found in sheep in this study. The absence of genomic group III genotypes in Spanish wildlife together with the wide distribution reported in domestic ruminants suggests that genotype

III+ may be a livestock genotype.

Genomic group IV has been detected in domestic - sheep and goat - and wild - rat and wild boar - animals, and in humans in a high number of Spanish provinces (Jado et al.,

2012; this study). The absence of genotypes in genomic group IV in wildlife in our study suggests that, although being able to infect wildlife, this group is mainly a livestock- human shared group of C. burnetii genotypes. Wildlife infected by genotypes in group

IV - wild boar and rat - are species with important links to human activities (Ruiz-Fons,

2015; Acevedo et al., 2014; Reusken et al., 2011; Meerburg & Reusken 2011) and may, therefore, get infected by interacting with livestock. In humans genomic group IV has been isolated from acute hepatitis cases, but is predominantly associated to chronic cases

(Jado et al., 2012).

Genomic group V has not been found in Spain to date including the results of this study.

A genotype (VI-) in the genomic group VI was found only in a red deer from southern

Spain in this study whereas a unique genotype (VII-) in genomic group VII was found in red deer and wild rabbits from south-central and southern Spain. Previous studies found

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C. burnetii genotypes in groups VI and VII in hard ticks of the species Hyalomma lusitanicum (VI and VII), Dermacentor marginatus (VII) and Rhipicephalus sanguineus

(VII) and also in Q fever human cases of acute hepatitis in southern and northern Spain, and in the Canary Islands (Jado et al., 2012). Interestingly, genotype VII- was, together with genotypes I+ and I-, the most frequent genotype found in wildlife in this study. This finding may suggest the existence of a wild cycle of these genomic groups between ticks and wild hosts - e.g. red deer and rabbits. Could a tick-wildlife cycle of C. burnetii be behind the geographic clustering of Q fever clinical presentations in humans in Spain?

Several epidemiologic evidences point to a potential implication of a tick-wildlife cycle with occassional transmission events to humans (directly through tick bites or indirectly through wildlife contaminated aerosols) in this geographic cluster of Q fever clinical presentation in humans: 1) The major part of C. burnetii genotypes isolated from patients with acute hepatitis in Spain belongs to genomic groups VI and VII (Jado et al., 2012);

2) Genotypes found in different tick species in central Spain belong to groups VI and VII;

3) These genomic groups are more prevalent (especially group VII) in red deer and wild rabbits from central and southern Spain; 4) Main red deer and rabbit distribution areas in

Spain are in central and southern Spain (see González-Barrio et al., 2015a, González-

Barrio et al., 2015b) where population densities of both species reach the highest reported values in Spain (Acevedo et al., 2008; Delibes-Mateos et al., 2008). Host density is a relevant factor driving the risk of infection by C. burnetii (Álvarez et al., 2012; Piñero et al., 2014; González-Barrio et al., 2015b); 5) Wild ungulates and small mammals are important hosts of Hy. lusitanicum and D. marginatus (Estrada-Peña et al., 2004; Ruiz-

Fons et al., 2006b; Ruiz-Fons et al., 2013). Whereas small mammals - e.g. rabbits - host immature stages of these tick species, wild ungulates host the adult stages (Hillyard et al.,

1996; Estrada-Peña et al., 2004; Ruiz-Fons et al., 2006b; Apanaskevich et al., 2008).

196

Domestic ungulates may also host these tick species although they’re frequently treated with antiparasitaries. Increasing density trends of deer in southern Spain (Acevedo et al.,

2008; Apollonio et al., 2010) promote increasing densities of ticks by increasing the reproductive success of adult ticks within the population (e.g. Ruiz-Fons & Gilbert,

2010b). Wild ungulates, especially red deer, are also important hosts for Rh. bursa, which as a member of the Rhipicephalus genus could also participate in a tick-wildlife cycle of

C. burnetii. Wild rabbits are important hosts to Rh. pusillus (Estrada-Peña et al., 2004), a tick in the Rh. sanguineus complex that has been also found positive to C. burnetii

(Toledo et al., 2009); 6) The map of the most suitable areas for Hy. lusitanicum in Spain

(Estrada-Peña et al., 2013) shapes the geographic distribution of acute hepatitis manifestations of Q fever in the country (Montejo et al., 1985; de Alarcón et al., 2003);

7) Red deer and domestic rabbits, and perhaps other wild hosts (González-Barrio et al.,

2015a, González-Barrio et al., 2015b, González-Barrio et al., 2015d) are reservoirs of C. burnetii in Spain; and 8) The 8.7% of questing Hy. lusitanicum ticks in central Spain are infected with C. burnetii (Toledo et al., 2009). Indeed, tick studies in northern Spain found a low prevalence of C. burnetii positive Haemaphysalis punctata whereas the most abundant exophilic tick species - Ixodes ricinus, H. inermis, H. concinna and D. reticulatus - were negative (Barandika et al., 2008). Red and roe deer are relevant hosts for adult stages of I. ricinus and H. concinna (Ruiz-Fons et al., 2006b). Additionally, both wild ungulate species have been found infected by C. burnetii in northern Spain (Ruiz-

Fons et al., 2008b; González-Barrio et al., 2015b). These eight points support the proposed hypothesis. However, current existing data on the host and geographic distribution of C. burnetii genotypes in Spain is not completely representative of the real status of the pathogen. Therefore, this hypothesis should be tested in future research

197 approaches and the information on circulating C. burnetii genotypes should be improved as well.

We didn’t find genomic group VIII in this study even though it has been reported in livestock and humans in mainland Spain and the Canary Islands (Jado et al., 2012).

Finally, the presence of the marker for virulence of C. burnetii genotypes (adaA) in wildlife also manifests the risks for humans of acquiring C. burnetii of wildlife origin

(van Schailk et al., 2013).

Conclusions

Coxiella burnetii genotypes are shared by wildlife, humans and livestock in Spain.

Certain C. burnetii genomic groups are more prone to be found in livestock whether others are more frequent in wildlife and ticks, suggesting particular pathogen-host adaptations that have been previously suggested (Capítulo IV). Molecular, epidemiological and ecological evidences suggest that wildlife - and certain tick species

- may be behind the geographic pattern observed in the clinical presentation of acute Q fever human cases in Spain.

Acknowledgements

We are grateful to colleagues at IREC (Jesús T. García, Javier Viñuela, Christian

Gortázar, Tania Carta, Mariana Boadella, José Ángel Barasona, João Queirós), to gamekeepers and to Francisco J. García for their help in wildlife surveys. We also acknowledge the collaboration of personnel at CERSYRA (Valdepeñas, Ciudad Real) to collect sheep samples and Rocío Jiménez Granado for kindky providing the goat bulk- tank milk samples. This study was funded by European Union FP7 ANTIGONE project

(278976) and partly by project RZ2010-00006-C02-01 of the Spanish Ministry for the

Economy and Competitiveness-MINECO. Grant support for this work was also from

INIA RTA2013-00051-C02-02 “Estudio de la viabilidad y caracterización de Coxiella

198 burnetii en explotaciones de pequeños rumiantes: dinámica y evolución de sus genotipos e implicaciones en Salud Pública”. F.R-F. acknowledges funding from MINECO through a ‘Ramón y Cajal’ research contract.

199

Capítulo III. Vías de

transmisión de Coxiella burnetii en fauna silvestre

ibérica.

200

Capítulo III. 1.

201

Vías de excreción de Coxiella burnetii y otros patógenos relevantes en

jabalí (Sus scrofa)

Shedding patterns of endemic Eurasian wild boar (Sus scrofa) pathogens

David González-Barrio, María Paz Martín-Hernando, Francisco Ruiz-Fons

Research in Veterinary Science. 2015. 102:206–211

202

Resumen

El jabalí ha experiementado un explosión demográfica en todo el mundo que aumenta el conocimiento sobre los patógenos compartidos. Sin embargo las rutas de excreción de los patógenos más relevantes en jabalí son desconocidas. Previas observaciones relacionadas con el sexo y la edad en el exposición del virus de la enfermedad de

Aujeszky (VEA) nos llevó a la hipótesis de que los patrones de excreción de patógenos endémicos de jabalíes pueden estar influenciados por los factores individuales. En este trabajo investigamos las rutas de excreción del virus de la enfermedad de Aujezky, el parvovirus porcino, el circovirus porcino tipo 2 y Coxiella burnetii y analizamos el efecto del sexo y la edad del hospedador en los patrones de excreción de los patógenos.

La presencia de anticuerpos de estos patógenos en suero fue analizada por medio de ensayo por inmunoadsorción ligado a enzimas (ELISA) y el AND del patógeno en hisopos orales, nasales, rectales y genitales fue analizado por medio de PCR. La influencia del sexo y la edad en la prevalencia de excreción de los patógenos fue analizada estadísticamente. Las principales rutas de excreción del virus de Aujeszky, del parvovirus porcino, del circovirus porcino tipo 2 y de Coxiella burnetii fueron identificadas, sin embargo la hipótesis en la relación del sexo y la edad con la excreción no se pudo confirmar.

203

Abstract

The Eurasian wild boar has experienced a worldwide demographic explosion that increases awareness on shared pathogens. However, shedding routes of relevant wild boar pathogens are unknown. Previous observations on sex- and age-related differences in

Aujeszky's disease virus (ADV) exposure led us to hypothesize that shedding patterns of endemic wild boar pathogens may be influenced by individual traits.We investigated shedding routes of ADV, porcine parvovirus (PPV), porcine circovirus type 2 (PCV2) and Coxiella burnetii and analysed the effect of host sex and age on pathogen shedding patterns. The presence of pathogen antibodies in serum and of pathogen DNA in oral, nasal, genital and rectal swabs was analysed by ELISA and PCR, respectively. The influence of sex and age in pathogen shedding prevalence was tested statistically. Main routes of ADV, PPV, PCV2 and C. burnetii shedding were identified but the hypothesis of sex- and/or age-related shedding patterns couldn't be confirmed.

204

Introduction

Populations of the Eurasian wild boar (Sus scrofa) have experienced an unprecedented demographic explosion in their native historic range in the last three decades, from

Western Europe to Japan (Cowled et al., 2009; Saito et al., 2012; Massei et al., 2015).

This demographic trend comes along with significant increase in pathogen prevalence and distribution (Gortázar et al., 2006; Ruiz-Fons et al., 2006, 2007; Meng et al., 2009), which is reflected by the spread of pathogens to naïve populations (Boadella et al., 2011;

EFSA, 2014) or by increasing prevalence of endemic pathogens (Vicente et al., 2005;

Boadella et al., 2012a; Pannwitz et al., 2012). These epidemiological change may increase the risk of transmission of shared pathogens at the wild boar-livestock human interface

(Ruiz-Fons et al., 2008). Pathogens such as Aujeszky's disease virus (ADV), porcine parvovirus (PPV), porcine circovirus type 2 (PCV2) and Coxiella burnetii may impact wild boar population dynamics (Ruiz-Fons et al., 2006; Schulze et al., 2010), threaten conservation of endangered species (Gortázar et al., 2010), interfere with pathogen control campaigns in domestic animals (Ruiz-Fons et al., 2008; Boadella et al., 2012b) or cause emerging disease outbreaks in humans (Tilburg et al., 2012). Understanding how and at which rate pathogens are transmitted both within wild boar and to in-contact species is essential to understand pathogen ecology and design effective preventive and control strategies (Boadella et al., 2012c). Aujeszky’s disease (AD), caused by suid alphaherpesvirus 1, affects both wild and domestic swine. Wild boar may constitute an important threat to ADV eradication campaigns or to the maintenance of AD-free status

(Boadella et al., 2012b). In pigs ADV is shed by nasal exudates, saliva, vaginal mucus, sperm, milk, faeces and occasionally urine. Similar, although not yet proven, shedding routes should be expected for wild boar. Some studies point to direct contact of wild boar as the most important pathway for ADV transmission (Romero et al., 2001; Ruiz-Fons et

205 al., 2007). On the basis of differing seroprevalence patterns between males and females, it was hypothesized (Ruiz-Fons et al., 2007) that oral/nasal transmission of ADV would constitute the main route of transmission between wild boar females within female groups, whereas transmission between sexes would be more frequent during the mating season (both by oral/nasal and venereal transmission). PPV causes reproductive failure in swine. Domestic pigs get infected by PPV by the oronasal, transplacental and veneral routes (Mengeling, 2006). However, transmission pathways in wild boar remain unknown in spite of the high PPV prevalence observed in free-roaming wild boar (Ruiz-Fons et al.,

2006; Roic et al., 2012). PCV2 is the aetiological agent of post-weaning multisystemic wasting syndrome in domestic pigs and wild boar (Schulze et al., 2003; Segalés et al.,

2012). The ability of PCV2 to infect bronchial and bronchiolar epithelial cells (Magar et al., 2000) and its isolation from bronchial, nasal, tonsilar, salivary, ocular, faecal and urinary swabs in domestic pigs (Segalés et al., 2005) suggest that oronasal secretions, urine and faeces are potential routes of viral shedding and transmission, also in wild boar.

Coxiella burnetii, the causal agent of Q fever, is a multi-host, highly environmentally resistant pathogen with high potential of causing human Q fever outbreaks (Tilburg et al.,

2012). Very recently, C. burnetii has been reported in Spanish wild boar (Astobiza et al.,

2011). However, nothing is known about infection and shedding pathways in suids to date. Shedding routes of C. burnetii in ruminants ˗ and perhaps also in suids ˗ are mainly vaginal secretions, faeces and milk (Maurin and Raoult, 1999). Our main goal was identifying shedding routes of ADV, PPV, PCV2 and C. burnetii in free-roaming wild boar. Previous observations on sex- and age-related differences in ADV exposure

(Romero et al., 2001; Ruiz-Fons et al., 2007) led us to hypothesize that shedding patterns of ADV, and perhaps of other endemic pathogens of wild boar, may be influenced by wild boar individual traits, e.g. sex and age.

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Materials and methods

Study area and sample collection

Hunter-harvested wild boar were surveyed in commercial hunting events (known as

“monterías”) during the 2010/2011 and 2012/2013 hunting seasons (from mid-October to mid-February) in 4 public hunting estates (ED, QM, RF and RO; Figure 1) located in

Montes de Toledo, south-central Spain (Vicente et al., 2005). Five individuals were surveyed in August. Animals were not particularly hunted for this study; the authors assisted to “monterías” for sample collection. We selected hunting estates that were identified as endemic for ADV, PPV and PCV2 (seroprevalences ranging 50%-60%) in previous studies (Vicente et al., 2004; Ruiz-Fons et al., 2006, 2007; Boadella et al.,

2012a). The status of C. burnetii was unknown, but the recent evidence of wild boar carrying C. burnetii in northern Spain (Astobiza et al., 2011) pointed out the need of investigating this zoonotic pathogen in our study populations. Sex was recorded and the age was estimated by tooth eruption patterns (Sáenz de Buruaga et al., 1991). Age was classified in three classes: i) juveniles (0-12 months old); ii) sub-adults (1-2 years old); and iii) adults (>2 years old). Nasal, oral, rectal and genital secretion samples were individually collected from surveyed wild boar with sterile swabs (Table 1) and preserved at 4ºC until arrival to the laboratory; genital swabs in males were collected by inserting the swab in the foreskin. Swabs were frozen at - 80ºC before four hours from collection.

Blood was collected from the cavernous sinus or from the thoracic cavity into sterile tubes and kept refrigerated until arrival to the laboratory. Blood was centrifuged at 3,000g for

10' and the serum preserved at -20ºC.

Serological analysis

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The presence of specific antibodies against ADV, PPV, PCV2 and C. burnetii in wild boar sera was analysed by commercial ELISA kits: IDEXX HerdCheck Anti-ADV gpl

(IDEXX Inc., USA; Ruiz-Fons et al., 2007), ELISA PPV compact (INGENASA, Spain;

Ruiz-Fons et al., 2006), ELISA CIRCO IgG (INGENASA, Spain; Boadella et al., 2012a) and LSI Q fever ruminant serum/milk ELISA kit (LSI, France; González-Barrio et al.,

2015), respectively.

Figure 1. Geographic location of the study hunting estates within peninsular Spain.

PCR analysis

DNA from swabs was extracted using a commercial kit (DNeasy ® Blood & Tissue kit,

Qiagen, Germany) following the protocols provided by the manufacturer. In order to improve DNA extraction from swabs, these were kept at 56ºC for 30' in a solution containing 20 μl of proteinase K and 200 μl of AL buffer, vortexed for 15" and discarded.

The sample remained at 56ºC for 30 additional minutes. After that, the manufacturer's blood extraction protocol was followed. DNA aliquots obtained were quantified

208

(NanoDrop 2000c/2000, Thermo Scientific, USA) and frozen at -20ºC until PCR performance. In order to discard sample cross-contamination, negative controls (Nuclease free water, Promega, USA) were included during DNA extraction. Every sample with a

DNA concentration over 50 ng/μl was homogenised to this concentration by adding the appropriate volume of sterile nuclease free water (Promega, USA). This step intended to optimize DNA concentration to be within the optimal range of the PCR mix employed

(PCR Master Mix, Promega, USA) according to the specifications of the manufacturer.

Later on, a nested conventional PCR targeting the highly conserved sequence of ADV glycoprotein B (gB) was performed on DNA from swabs (oral, nasal, rectal and genital) as previously described (Ruiz-Fons et al., 2007). Detection of PPV and PCV2 DNA in swabs was carried out by a multiplex conventional PCR targeting the NS1 gene of PPV and the ORF1 gene of PCV2 as described elsewhere (Jiang et al., 2010). Finally, swabs were analysed by means of a conventional PCR targeting a transposon-like repetitive region of C. burnetii as previously described (Astobiza et al., 2011). PCR products were elicited by PCR fragment size estimation in comparison with molecular weight standard

'GeneRuler 100 bp Plus DNA Ladder' (Thermo Scientific, USA) after electrophoresis on

1.2% agarose gel containing 0.1 μl/ml GelRedTM Nucleic Acid Gel Stain (Biotium,

USA). DNA extraction and PCR were performed in separate laboratories under biosafety level II conditions (BIO II A Cabinet, TELSTAR, Spain) to avoid cross-contamination.

PCR primer details are shown in Table 1.

Data analysis

Pathogen shedding by wild boar was estimated on the basis of the presence/absence of pathogen DNA in swabs. Chi-square tests of homogeneity - or Fisher's exact tests when

209 required (expected cell frequency < 5) - were employed to assess for the relationship between pathogen shedding prevalence - both overall (PCR positive in either oral, nasal or genital swabs) and route-specific shedding (nasal, oral or genital in separate) - and host individual traits (sex and age class). IBM SPSS 22.0 (IBM, Armonk, NY, USA) was employed for statistical analyses. The statistical uncertainty for each of the estimated proportions was assessed by calculating the associated Clopper-Pearson exact 95% confidence interval (95%CI) with Quantitative Parasitology 3.0

(http://www.zoologia.hu/qp/qp.html).

Table 1. Reported shedding routes of Aujeszky’s disease virus (ADV), porcine parvovirus (PPV), porcine circovirus type 2 (PCV2) and Coxiella burnetii in domestic animals, detailed information on swabs collected in wild boar to study shedding routes and sequences of primers employed for PCR analyses.

Pathogen Shedding routes Collected swabs Primer ref. Sequence (5´-3´) Target products (bp) ADV Nasal, oral, genital and Nasal, oral, rectal, fADVgB1a ATGGCCATCTCGCGGTGC gB gene (334)e milka genital ADVgB1b ACTCGCGGTCCTCCAGCA

fADVgB2a ACGGCACGGGCGTGATC gB gene (195)e

ADVgB2b GGTTCAGGGTACCCCGC

PPV Nasal, oral and genitalb Nasal, oral, rectal, PVF AGTTAGAATAGGATGCGAGGAA NS1 gene (265)f genital PVR AGAGTCTGTTGGTGTATTTATTGG

PCV2 Nasal, oral, rectal and Nasal, oral, rectal, CVF CGAGAAAGCGAAAGGAACAGA ORF 1 (371)f genitalc genital CVR GGTAACCATCCCACCACTT Coxiella burnetii Rectal, genital and Nasal, oral, rectal, Trans1 TATGTATCCACCGTAGCCAGTC Transposon-like milk in ruminantsd; genital repetitive region (687)g unknown in suids Trans2 CCCAACAACACCTCCTTATTC aPejsak and Truszczynski, 2006; bMengeling, 2006; cSegalés et al., 2005; dMaurin and Raoult,

1999; eBalasch et al., 1998; fJiang et al., 2010; gBerri et al., 2000

Results

In total, 133 wild boar − 68 males and 65 females − that corresponded to 22 juveniles (9 males and 13 females), 29 sub-adults (18 males and 11 females) and 82 adults (41 males and 41 females) were surveyed. ELISA results for ADV, PPV, PCV2 and C. burnetii are shown in Table 2. ADV DNA was detected in oral, rectal and genital secretions but not

210 in nasal secretions; DNA in oral and rectal secretions was detected only in females whereas positive genital secretions were detected both in males and females (Table 3).

ADV DNA was only detected in swabs from juveniles (1/18) and adults (7/67); only a rectal swab was positive in juveniles whereas oral and genital swabs were positive in adults (Table 4). PPV DNA was detected in nasal, rectal and genital swabs, but not in oral swabs. In both sexes PPV DNA was detected in nasal and rectal swabs, but positive genital swabs were only detected in males (Table 3). PPV positive swabs were mainly detected in adults (Table 4). PCV2 DNA was detected in nasal, oral and rectal secretions but not in genital secretions; oral and nasal swabs from males and females contained

PCV2 DNA whereas only rectal swabs from females were PCR positive (Table 3). PCV2

PCR positive swabs were detected in juveniles, sub-adults and adults (Table 4). Coxiella burnetii DNA was detected in nasal, rectal and genital swabs but not in oral swabs.

Positive nasal secretions were evidenced in both males and females, but only females were positive in rectal secretions and only males in genital secretions (Table 3). One of

17 juveniles and 1 of 26 sub-adults had C. burnetii DNA in nasal secretions whereas 1 of

63 and 1 of 66 adults presented C. burnetii DNA in rectal and genital swabs, respectively

(Table 4). No statistically significant relationship was evidenced between host individual traits and pathogen shedding prevalence in any of the analysed pathogens.

Discussion

This paper explores the shedding routes of endemic pathogens in wild boar by molecular studies while current knowledge on wild boar pathogens has been mainly based on serological surveys. This study tested the hypothesis that host individual factors such as sex and age, presumably linked to behaviour or physiology (Ruiz-Fons et al., 2013), would modulate pathogen shedding patterns in Eurasian wild boar, therefore exerting variation to the role of individuals in pathogen transmission. This hypothesis couldn’t be

211 confirmed, perhaps because of the low shedding prevalence of ADV, PPV, PCV2 and C. burnetii in the studied populations.

Table 2. Positive samples in ELISA over sampling size (Pos/N), and seroprevalence (in %) and associated exact 95% confidence interval (within brackets), of Aujeszky´s disease virus (ADV), porcine parvovirus (PPV), porcine cirvovirus type 2 (PCV) and Coxiella burnetii throughout wild boar sex and age. Sex Age ADV PPV PCV2 Coxiella burnetii

Male Juvenile 4/7 2/8 3/7 0/8 (57.1; 18.4-90.1) (25.0; 3.2-65.1) (42.9; 9.9-81.6) (0.0; 0.0-37.0) Sub-adult 8/16 6/16 11/16 0/16 (50.0; 24.7-75.4) (37.5; 15.2-64.6) (68.8; 41.3-90.0) (0.0; 0.0-20.6) Adult 18/38 28/38 28/38 1/38 (47.4; 31.0-64.2) (73.7; 56.9-86.6) (73.7; 56.9-86.6) (2.6; 0.1-13.8) Total male 30/61 36/62 42/61 1/62 (49.2; 36.1-62.3) (58.1; 44.8-70.5) (68.9; 55.7-80.1) (1.6; 0.0-8.7) Female Juvenile 10/12 3/12 8/12 0/12 (83.3; 51.6-97.9) (25.0; 5.5-57.2) (66.7; 34.9-90.1) (0.0; 0.0--26.5) Sub-adult 7/10 4/10 8/10 0/10 (70.0; 34.8-93.3) (40.0; 12.2-73.8) (80.0; 44.4-97.5) (0.0; 0.0-30.9) Adult 23/41 29/41 35/41 0/41 (56.1; 39.7-71.5) (70.7; 54.5-83.9) (85.4; 70.8-94.4) (0.0; 0.0-8.6) Total female 40/63 36/63 51/63 0/63 (63.5; 50.4-75.3) (57.1; 44.0-69.6) (81.0; 69.1-89.8) (0.0; 0.0-5.7) Total 70/124 72/125 93/124 1/125 (56.0; 47.3-65.3) (57.6; 48.4-66.4) (75.0; 66.4-82.3) (0.8; 0.0-4.4)

Table 3. Samples positive by PCR in relation to sample size (Aujeszky’s disease virus (ADV), porcine parvovirus (PPV), porcine circovirus type 2 (PCV2) and Coxiella burnetii) in secretions according to sex. Prevalence (in %) and associated 95% exact confidence interval are shown within brackets. Detection of positive samples is bold marked. Swab Pathogen Sex Nasal Oral Rectal Genital ADV Male 0/62 0/62 0/59 5/60

(0.0; 0.0-5.8) (0.0; 0.0-5.8) (0.0; 0.0-6.1) (8.3; 2.8-18.4) Female 0/54 1/53 1/49 1/50

(0.0; 0.0-6.6) (1.9; 0.0-10.1) (2.0; 0.0-10.9) (2.0; 0.1-10.7) Total ADV 0/116 1/115 1/108 6/110

(0.0; 0.0-3.1) (0.9; 0.0-4.8) (0.9; 0.0-5.1) (5.5; 2.0-11.5) PPV Male 3/63 0/62 2/56 1/59

(4.8; 1.0-13.3) (0.0; 0.0-5.8) (3.6; 0.4-12.3) (1.7; 0.0-9.1) Female 1/52 0/49 2/46 0/47

(1.9; 0.0-10.3) (0.0; 0.0-7.3) (4.3; 0.5-14.8) (0.0; 0.0-7.6) Total PPV 4/115 0/111 4/102 1/106

(3.5; 1.0-8.7) (0.0; 0.0-3.2) (3.9; 1.1-9.7) (0.9; 0.0-5.2) PCV2 Male 2/62 2/61 0/56 0/59

(3.2; 0.4-11.2) (3.3; 0.4-11.4) (0.0; 0.0-6.4) (0.0; 0.0-6.1) Female 1/52 1/51 1/47 0/47

(1.9; 0.0-10.3) (2.0; 0.0-10.5) (2.1; 0.0-11.3) (0.0; 0.0-7.6) Total PCV2 3/114 3/112 1/103 0/106

(2.6; 0.5-7.5) (2.7; 0.6-7.6) (1.0; 0.0-5.3) (0.0; 0.0-3.4) Coxiella Male 1/62 0/61 0/54 1/58

burnetii (1.6; 0.0-8.7) (0.0; 0.0-5.9) (0.0; 0.0-6.6) (1.7; 0.0-9.2) Female 1/50 0/50 1/46 0/47

(1.9; 0.0-10.7) (0.0; 0.0-7.1) (2.2; 0.0-11.5) (0.0; 0.0-7.6) 212 Total C. burnetii 2/112 0/111 1/100 1/105

(1.8; 0.2-6.3) (0.0; 0.0-3.2) (1.0; 0.0-5.5) (1.0; 0.0-5.2)

Methodological considerations

Surveying hunter-harvested wild boar gives access to a high number of individuals but tissue damage by bullets or hunting dogs may limit the collection of specific samples.

Tissue damage precluded collecting nasal, oral, rectal and genital swabs from every surveyed individual. In order to test the effect of host traits in shedding prevalence patterns, sample size was estimated with described seroprevalence rates of endemic pathogens. However, the low shedding prevalence found may have perhaps impaired the robustness of statistical findings with selected sample sizes for this study. Prevalence of

ADV, PPV and PCV2 DNA in secretions was lower than expected according to observed seroprevalences. Could sampling bias to the hunting season be responsible for the low shedding prevalence observed? Whether pathogen shedding is seasonal or not in wild boar is currently unknown. Seasonal shedding would be expected for pathogens shed mainly around the breeding season ˗ e.g. C. burnetii (Maurin and Raoult, 1999) and PPV

(Mengeling, 2006) ˗ that in Spanish wild boar takes place around the end of January to

February (Vicente et al., 2005). Thirty-two of the 133 wild boar (24.1%) were surveyed in January-February whereas 101 wild boar were surveyed from August to December.

Overall prevalence in swabs ˗ excluding results from rectal swabs ˗ was similar in Jan-

Feb (1/23 ˗ 4.3%; 95% C.I.: 0.1-22.0) than in Aug-Dec (3/76 ˗ 3.9%; 95%C.I.: 0.8-11.1) for C. burnetii DNA, ADV DNA (2/25 ˗ 8.0%; 95% C.I.: 1.0-26.0 ˗ and 6/84 ˗ 7.1%; 95%

226 C.I.: 2.7-14.9, respectively), PPV DNA (1/22 ˗ 4.5%; 95% C.I.: 0.1-22.9 ˗ and 3/72

- 4.2%; 95% C.I.: 0.9-11.7, respectively) and PCV2 DNA (1/22 ˗ 4.5%; 95%C.I.: 0.1-

22.9 ˗ and 4/73 ˗ 5.5%; 95%C.I.: 1.5-13.4, respectively). Therefore, sampling wild boar during the hunting season, which covers from mating to breeding seasons, seems adequate for studying shedding routes and prevalence of these pathogens. Although detection of pathogen genetic material does not necessarily reflect pathogen viability, we herein

213 assumed that pathogen DNA detection in nasal, oral and genital swabs and tissues reflected the presence of viable pathogen whereas detecting pathogen DNA in rectal swabs did not since pathogen DNA could come from ingestion of infected wild boar tissues (cannibalism) or through ingestion of secretions.

Table 4. Samples positive by PCR over sampling size (Aujeszky’s disease virus (ADV), porcine parvovirus (PPV), porcine circovirus type 2 (PCV2) and Coxiella burnetii) in secretions throughout wild boar age. Prevalence (in %) and associated 95% exact confidence interval are shown within brackets. Positive samples detected are bold marked.

Pathogen shedding patterns

Pathogen shedding prevalence - both overall and route-specific shedding - was not statistically influenced by sex and age, and therefore our initial hypothesis couldn’t be

214 confirmed for any of the studied pathogens. The lack of statistical significance was most probably linked to the limited sample size for the low shedding prevalence found.

Therefore, our findings will be discussed in terms of average shedding prevalence. The predominant shedding of ADV in genital secretions suggests that the main route for ADV transmission in Eurasian wild boar is venereal. This finding agrees with previous observations in Italy (Verin et al. 2014) and in the US (Romero et al., 2001). However, shedding of ADV in oral secretions by females would suggest that oral transmission occurs within female groups. In this study none of the nasal swabs analysed had ADV

DNA, but Verin et al. (2014) found nasal shedding of ADV in naturally infected wild boar. Altogether, these results shape what we’d expect from the initial hypothesis of predominant venereal transmission between males and females (Ruiz-Fons et al., 2007) at the time of mating (as previously suggested by Vicente et al., 2005) whereas ADV would be transmitted oro-nasally within female groups along the year. PPV seroprevalence was similar in both sexes, suggesting that males and females are equally exposed to infection by PPV. This could be due to the high probability of indirect transmission through contaminated fomites caused by the high environmental resistance of PPV (Mengeling, 2006). Observed shedding prevalence suggests a predominance of nasal over venereal transmission of PPV in wild boar. The oro-nasal route seems to be predominant for PCV2 transmission in wild boar because of the higher shedding prevalence in nasal and oral than in genital secretions. This observation would fit our expectations since it is known that PCV2 is transmitted mainly by oronasal secretions in domestic swine (Segalés et al., 2005). Coxiella burnetii is becoming an increasing concern for both animal and public health authorities (Tilburg et al., 2012). The demographic explosion of wild boar in Europe may influence C. burnetii ecology and this was the main reason for analysing the status of C. burnetii in highly dense and aggregated

215 wild boar populations (Boadella et al., 2011; Ruiz-Fons et al., 2007). Coxiella burnetii prevalence exceeds seroprevalence in wild ungulates (Rijks et al., 2011), and this pattern

- which perhaps relays on the fact that a fraction of infected animals do not seroconvert

(De Cremoux et al., 2012) - was also evidenced in wild boar. We found C. burnetii DNA in wild boar male genital secretions, which suggests that C. burnetii may be transmitted venereally in wild boar. Since artificial insemination is a frequent practice in industrialized countries, the risk of venereal transmission by males should not be dismissed (see Ruiz-Fons et al., 2014).

Conclusions

In this work we identified routes of ADV, PPV, PCV2 and C. burnetii transmission between wild boar and potentially to in-contact third species. The initial hypothesis of an effect of wild boar individual traits on shedding patterns, and therefore on potential transmission patterns, couldn’t be confirmed. This was perhaps linked to the low shedding prevalence found in spite of ˗ or perhaps linked to ˗ the high ADV, PPV and PCV2 seroprevalence in the studied populations. Host population-pathogen interaction traits such as population (herd) immunity may also modulate shedding prevalence and routes for certain wild boar endemic pathogens (see Piñero et al., 2014), indicating that efficient prevention and control of these pathogens would only be achieved if their dynamics is properly understood.

Acknowledgements

We acknowledge collaboration from authorities (Junta de Comunidades de Castilla ˗ La

Mancha, Sociedad de Cazadores Estados del Duque de Malagón and Organismo

Autónomo de Parques Nacionales OAPN) for sample collection. This study is a contribution to European Commission grants ANTIGONE (278976) and APHAEA

216

(EMIDA ERA-NET) and to agreement between OAPN and IREC for “Quintos de Mora” estate. FRF acknowledges funding from “Juan de la Cierva” and “Ramón y Cajal” contracts (Spanish Ministry for the Economy and Competitiveness - MINECO).

217

Capítulo III. 2.

218

Vías de excreción de Coxiella burnetii en ciervo rojo (Cervus elaphus)

en condiciones de producción semi-extensiva.

Coxiella burnetii shedding by farmed Red Deer (Cervus elaphus)

David González-Barrio, Sonia Almería, María Rosa Caro, Jesús Salinas, José Antonio

Ortiz, Christian Gortázar and Francisco Ruiz-Fons.

Transboundary and Emerging Diseases. 2015. 62: 572–574

219

Resumen

La fauna silvestre, y en concreto algunas especies de ciervo, debido al importante incremento mundial de granjas de ciervo, pueden contribuir al mantenimiento de Coxiella burnetii, el agente causante de la fiebre Q. Actualmente, no existen precedentes que vinculan la exposición a especies de ciervos con los casos de fiebre Q humana. Sin embargo, un caso de fiebre Q en humano fue diagnóstica recientemente en una granja de ciervos (Cervus elaphus), y que nos llevó a investigar si los ciervos podrían ser una fuente de contaminación del medio ambiente con C. burnetii y determinar la implicación de C. burnetii en el fracaso reproductivo en la granja. Sueros sanguíneo e hisopos vaginales fueron cogidos de ciervas con y sin fallo reproductivo, y testado para detectar la presencia de anticuerpos y ADN de Coxiella burnetii, Chlamydia abortus, Neospora caninum y

Toxoplasma gondii. Los resultados de las serologías y de las PCRs sugieren que Coxiella burnetii fue la primera causa del fallo reproductivo. De esta manera identificamos la excreción vaginal de Coxiella burnetii por parte de las ciervas, confirmando así al ciervo como una fuente de infección por la enfermedad zoonótica fiebre Q

220

Summary

Wildlife and notably deer species – due to the increasing relevance of deer farming worldwide – may contribute to the maintenance of Coxiella burnetii, the causal agent of

Q fever. Currently, there are no precedents linking exposure to deer species with human

Q fever cases. However, a human case of Q fever was recently diagnosed in a red deer

(Cervus elaphus) farm, which led us to investigate whether deer could be a source for environmental contamination with C. burnetii and ascertain the implication of C. burnetii in reproductive failure in the farm. Blood serum and vaginal swabs were collected from hinds either experiencing or not reproductive failure and tested to detect the presence of antibodies and DNA, respectively, of C. burnetii, Chlamydia abortus, Neospora caninum and Toxoplasma gondii. Serology and PCR results suggest C. burnetii was the primary cause of the reproductive failure. We identified vaginal shedding of C. burnetii in hinds, confirming red deer as a source of Q fever zoonotic infection.

221

Introduction

Q fever is a worldwide zoonosis caused by Coxiella burnetii, an obligate intracellular bacterium. This bacterium is an important cause of reproductive failure in domestic ruminants -cattle, sheep, goats- and other mammals (Maurin and Raoult, 1999). The major part of human Q fever outbreaks are linked to domestic ruminants (e.g. Tilburg et al., 2012), which shed high loads of C. burnetii to the environment around the breeding season. Coxiella burnetii can be thereafter disseminated through aerosols that constitute the main pathway for human infection. Wildlife and notably deer species can also contribute to the maintenance of this multihost pathogen (Ruiz-Fons et al., 2008).

Exposure to wild deer, for instance of hunters during, game carcass dressing, has been proposed as a potential zoonotic risk (Kirchgessner et al., 2012). However, there are no precedents linking exposure to deer species with human Q fever cases. Deer farming is important in regions such as New Zealand, Europe and North America, but there is no published evidence of reproductive failure in deer due to infection with C. burnetii.

This scenario emphasizes the need for a better understanding of the role of C. burnetii in the reproductive failures in farmed deer. Our goal was investigating the implication of C. burnetii in reproductive failure in farmed red deer (Cervus elaphus) to ascertain whether deer could be a source of C. burnetii in a farm with a history of a human case of Q fever.

This case, an acute infection requiring hospitalization, affected the veterinarian in charge of the deer. This person had no professional exposure to other domestic ruminants. On the basis of previous results on red deer and C. burnetii showing 29% serum antibody prevalence and a wide spatial distribution of seropositive deer in Spain (Ruiz-Fons et al.,

2008), we hypothesized that C. burnetii might cause reproductive failure in deer farms and constitute a zoonotic risk.

222

Materials and Methods

The study was performed on a semi-extensive red deer farm in southern Spain. The number of deer in the farm was 410 hinds and 72 stags. Reproductive failure had been documented by the farm veterinarian over the last 10 years. In spring 2011, 27 of 350 echography-confirmed pregnant hinds (7.7%) failed to carry a calf. In September 2011, 5 months after the calving season and during weaning, blood samples were collected from

12 hinds with reproductive failure and from 13 hinds that calved normally. Vaginal swabs were taken from 10 hinds with reproductive failure and 13 that calved normally.

Blood serum was tested for antibodies against C. burnetii by LSI Q fever ruminant serum/milk ELISA kit (Life Technologies, Grand Island, NY, USA). Sera were also tested against other known causes of reproductive failure including Clamydia abortus (in- house blocking ELISA using a recombinant polymorphic outer membrane protein as antigen; Salinas et al., 2009), Neospora caninum (competitive ELISA; VMRD, Pullman,

WA, USA) and Toxoplasma gondii (modified agglutination test, MAT; Dubey and

Desmonts, 1987).

For PCR, DNA was extracted from vaginal swabs using the DNeasy Blood and Tissue kit (QIAGEN, Hilden, Germany). The C. burnetii htpB gene was amplified by nested

PCR as previously reported (To et al., 1996). A PCR targeting the pmp 90 of 91 gene was performed to detect C. abortus (Salinas et al., 2012). For N. caninum, the specific genomic Nc5 region was targeted and the PCR performed as previously described

(Darwich et al., 2012). Finally, for T. gondii, a nested PCR for detection of the 529-bp repetitive fragment was performed (Darwich et al., 2012).

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Differences in seroprevalence and DNA prevalence between hinds with and without reproductive failure were assessed by means of homogeneity tests. Statistical uncertainty linked to sample size was assessed for each prevalence by calculating the 95% confidence interval (CI) according to the expression 95% CI = 1.96 [p(1–p)/n]1/2, where ‘p’ is the unitary value of the proportion and ‘n’ is the sample size.

Results and Discussion

The overall seroprevalence of C. burnetii was 36% (95% CI: 17.2–54.8). The seroprevalence in hinds with reproductive failure was 50% (95% CI: 21.7–78.3), while seroprevalence in hinds that calved normally was 23.1% (95% CI: 0.0–46.0). However, this difference was not statistically significant (P > 0.05). In the case of C. abortus, the overall seroprevalence was 32% (95% CI: 13.7–50.3), with similar antibody prevalence among hinds with (33.3%; 95% CI: 6.6–60.0) and without (30.8%; 95% CI: 5.8–55.8) reproductive failure (P > 0.05). By contrast, all samples were seronegative to N. caninum, and seroprevalence to T. gondii was low, 8% (95% CI: -3.4 to 18.6), with similar seroprevalence observed in hinds with (8.3%; 95% CI: -7.3 to 23.9) and without (7.7%;

95% CI: -6.8 to 22.2) reproductive failure (P > 0.05).

Regarding the detection of specific DNA, C. burnetii DNA was detected in 26.9% (95%

CI: 8.8–45.0) of the vaginal swabs. The prevalence differed between hinds with (40%;

95% CI: 9.7–70.3) and without (15.4%; 95% CI: -4.2 to 35.0) reproductive failure (P >

0.05). By contrast, none of the vaginal swabs analysed contained DNA of N. caninum, T. gondii and C. abortus.

The DNA results, together with the higher C. burnetii seropositivity in hinds with reproductive failure, as well as the lack of associations with the other pathogens analysed, suggest that Q fever could have been the primary cause of the reproductive failure

224 outbreak in the deer farm. A crosssectional survey on C. burnetii in deer in Spain found that 40% of red deer from this particular farm (both adults and juveniles) had been exposed to C. burnetii (Ruiz-Fons et al., 2008). This, along with the current results, suggests that the pathogen is endemic in this farm. For comparison, the average herd serum antibody prevalence of C. burnetii in small ruminants in endemic areas of northern

Spain did not exceed 15%. Production systems (more intensive vs. extensive) were suspected to play a relevant role in exposure to C. burnetii (Ruiz-Fons et al., 2010).

Excretion routes of C. burnetii in wild ungulates are unknown, and shedding patterns remain to be determined. However, abortive materials and vaginal excretion are likely sources of contamination for in-contact persons, such as deer-farm personnel. In this study, C. burnetii DNA was still present in vaginal swabs 5 months after calving. This indicates that C. burnetii was shed in vaginal mucus for a long period after parturition or reproductive failure, constituting a source for environmental contamination and zoonotic

Q fever. Further studies are needed to fully elucidate the epidemiology of C. burnetii in farmed and wild deer, particularly regarding excretion routes and transmission risks.

Acknowledgements

We are grateful to farm keepers for their help with the survey. This work was funded by

EU FP7 Grant ANTIGONE (278976) and CDTI (Centro para el Desarrollo Tecnologico

Industrial, Spanish Ministry for Economy and Competitiveness). F. Ruiz-Fons is supported by a Juan de la Cierva contract from the Spanish Ministry for Economy and

Competitiveness.

225

Capítulo IV

226

Estrategias de control de Coxiella burnetii: evaluación de la vacunación

con vacunas inactivadas comerciales de fase I como estrategia de

reducción de la prevalencia y el nivel de excreción de la bacteria en

ciervo rojo (Cervus elaphus)

Evaluating the efficiency of commercial phase I inactivated Coxiella burnetii

vaccine in decreasing infection prevalence and shedding in red deer (Cervus

elaphus)

David González-Barrio, Jose Antonio Ortiz, Francisco Ruiz-Fons

En preparación

227

Resumen

El papel de la fauna silvestre está incrementando como una pieza relevante en el ciclo de vida de Coxiella burnetii. El ciervo rojo puede ser uno de los más relevantes reservorios silvestre de C. burnetii en Europa donde sus poblaciones muestran un notable incremento en su demografía y en sus tendencias de distribución geográficas. La evaluación de la eficacia de las herramientas para el control de la infección por C. burnetii en ciervos rojos podría ser esencial para prevenir eventos de transmisión de la fauna silvestre-ganado- humanos. Se diseñó un programa de vacunación en una población de ciervos rojos en sistema de producción semi-extensiva utilizando una vacuna inactivada comercial frente a C. burnetii. Las ciervas adultadas y jóvenes de un año se vacunaron, y una dosis de recuerdo se administró 3 semanas depués de la primera vacunación. Posteriormente se administró una vacuna annual y biannual. La eficacia de la vacuna se midió en términos de su potencial en la reducción de la prevalencia y en la disminución de la carga de excreción de C. burnetii. La excreción en secreciones vaginales, leche y heces fue evaluada in ciervas vacunadas y en el grupo control después del parto a lo largo de tres años desde el inicio del programa de vacunación. La respuesta immune humoral a la vacunación y a la infección fue seguida en la población de studio desde antes del inico de la vacunación hasta 3 años después. Aunque la vacunación en las ciervas indujo altas tasas de seroconversion, especialmente en los animales adultos, no hubo una reducción en la excreción de C. burnetii en las secreciones vaginales y la leche. Sin embargo, la excreción en las heces experiment una reducción notable a lo largo del período de studio después de la vacunación, tanto en animales vacunados como en los animales coexistentes del grupo control. Este hallazgo, además de lo encontrado previamente en la reducción de la presión de la infección sobre los animales no tratados previamente en la granja desde

228 el inicio de la vacunación sugiere una possible eficacia de la vacunación en ciervos para la reducción de la contaminación ambiental por C. burnetii.

229

Abstract

Wildlife is increasingly being faced as a relevant piece in the life-cycle of Coxiella burnetii. Red deer may be one of the most relevant wild reservoirs for C. burnetii in

Europe where its populations display notable increasing demographic and geographic distribution trends. Evaluating the effectiveness of tools to control infection by C. burnetii in red deer could be essential to prevent wildlife-livestock-human transmission events.

We designed a vaccination program in a semi-extensively bred red deer population by using a commercial phase I inactivated C. burnetii vaccine. Adult and yearling deer females were vaccinated and a boost dose was given 3 weeks later. Annual and biannual revaccination was applied subsequently. The efficiency of the vaccine was measured in terms of its potential to reduce C. burnetii shedding prevalence and burden. Shedding in vaginal secretions, milk and faeces was evaluated in vaccinated deer and in control groups after calving along three years from the beginning of the vaccination program. The humoral immune response to vaccination and to natural infection was followed in the study population from before the start of vaccination up to 3 years later. Although vaccination of deer females induced high seroconversion rates, especially in adult animals, there was not a reduction in C. burnetii shedding in vaginal secretions and milk.

However, shedding in faeces experienced a notable reduction along the study period after vaccination both in vaccinated animals and in coexisting mates of the control group. This finding in addition to previous findings of a reduction in infection pressure over naïve animals in the farm from the onset of vaccination point to a potential efficiency of deer vaccination in reducing environmental contamination by C. burnetii. Results suggest that faeces could constitute the main source for environmental contamination with C. burnetii.

Further experiments should be carried out in the future to test different vaccination approaches and monitor the effect of vaccination over a longer period of time.

230

Introduction

Coxiella burnetii causes Q fever, a worldwide distributed zoonotic infectious disease.

Humans are dead-end hosts that become infected by the inhalation of infected aerosols that carry over infectious C. burnetii bacteria shed by infected animals (Maurin and

Raoult, 1999). Livestock - mainly ruminant species such as cattle, sheep and goats - are the main source of C. burnetii infected aerosols that can infect humans. Nonetheless, recent scientific evidences also point to wildlife as a relevant piece in the life cycle of C. burnetii and therefore as a potential source of C. burnetii for livestock and humans

(Gonzalez-Barrio et al., 2015a,b,c). Coxiella burnetii is widely distributed among livestock and wildlife populations in the World. This fact confers human Q fever a global public health relevance that makes worthy investing resources on investigating potential control tools at origin, in the animal reservoir.

Infection by C. burnetii in animals - mainly in cattle, sheep and goats (Agerholm 2013) but also in wildlife (Clemente et al., 2008; González-Barrio et al., 2015 TED) - has been associated with sporadic cases of stillbirth, premature delivery, abortion and weak offspring. The clinical outcome of Q fever in livestock carries over significant productive losses (Oporto et al., 2006; García-Ispierto et al., 2014) and in wildlife it also may constitute an important drawback in the conservation of endangered ruminant species in zoological collections (Teresa Albaigar, personal communication). Infected animal females shed high burdens of C. burnetii into the environment around parturition after normal delivery or after reproductive failure. Coxiella burnetii is mainly shed in vaginal secretions, milk and faeces (Angelakis and Raoult, 2010; Astobiza et al., 2011). These secretions, mainly vaginal mucus and faeces, contaminate the environment from which

C. burnetii may be transmitted to susceptible hosts.

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The relevance of Q fever in animal and public health makes essential the promotion of research on potential prevention and control tools in livestock and wildlife. Any measure leading to a decrease in the prevalence of shedders and in the burden of shed bacteria would reduce environmental contamination and would limit the spread of the infection.

The optimal strategy for controlling infection by C. burnetii in livestock is a combination of measures, being vaccination one of the most effective ones (Arricau-Bouvery et al.,

2005; EFSA 2010). Vaccines composed of antigenic phase I bacteria award higher protection than those prepared with phase II C. burnetii (Arricau-Bouvery et al., 2005).

The largest known vaccination campaign in small ruminants with an inactivated phase I vaccine started at the end of 2008 in the Netherlands, a year after the beginning of the massive 2007-2010 outbreak of human Q fever (van de Brom et al., 2009; Schimmer et al., 2011; Roest et al., 2011). The vaccination campaign of domestic ruminants together with additional measures reduced the incidence of Q fever in humans to normal levels

(Vellema et al., 2014). The efficacy of commercial phase I inactivated vaccines has been assessed in naturally infected cattle, goat and sheep populations (Guatteo et al., 2008;

EFSA 2010; Hogerwerf et al., 2011; Rousset et al., 2009; Astobiza et al., 2011), demonstrating protection in non-infected susceptible animals. The use of an inactivated phase I vaccine in previously non-infected goats and cattle was associated with a strong reduction in the incidence of abortion, reduction of vaginal shedding and disappearance of C. burnetii shedding in milk (Arricau-Bouvery et al., 2005; Guatteo et al., 2008). These results indicate that the use of inactivated phase I vaccines may be an effective method to reduce the risk of infection by C. burnetii when administered to uninfected or low prevalence populations. The efficiency of these vaccines in infected populations is questionable (Astobiza et al., 2011a; Guatteo et al., 2008), especially if vaccination is carried out during a short period (˂ 1 year) of time. However, in the real world, C. burnetii

232 is endemic in a high number of ruminant populations (Ruiz-Fons et al., 2010; Saegerman et al., 2015; González-Barrio et al., 2015 AEM y EID). This fact makes necessary to estimate the potential efficiency of commercially available phase I vaccines to control C. burnetii infection in endemic animal populations.

Currently, no trial has evaluated the efficiency of inactivated phase I C. burnetii vaccines in any wild species, either free-roaming or in captivity. However, any new study aiming to determine the status of C. burnetii in wildlife shows wide geographic and host ranges.

Furthermore, C. burnetii circulates in wildlife populations at similar or higher prevalence than in livestock populations. In Iberia, several (50%) red deer (Cervus elaphus) populations are endemically infected by C. burnetii, which suggest that red deer may be a relevant wild reservoir of C. burnetii (González-Barrio et al., 2015 AEM). Red deer is one of the main big game species in Europe, which makes its populations the subject of introduction of management practices that promote hunting (Vicente et al., 2006). Red deer farming has also expanded worldwide in recent decades due to the demand of venison and also caused by population restocking with live farm-bred and genetically controlled individuals (Hoffman & Wicklund, 2008; Griffiths et al., 2010). These facts support the need of research in potential C. burnetii control tools in red deer. Indeed, this was the aim of this study in which an experimental field vaccination trial with a commercial inactivated phase I vaccine was designed for a C. burnetii endemic red deer semi-extensively bred population. The efficiency of the vaccination experiment was evaluated in terms of reduction in C. burnetii shedding prevalence and burden along a 3- year period post-vaccination.

233

Materials and methods

The red deer study population

The study was performed in a semi-extensively bred red deer population located in the province of Cádiz (Southern Spain) in which C. burnetii circulates endemically (Ruiz-

Fons et al., 2008; González-Barrio et al., 2015a; González-Barrio et al., 2015d). Deer are semi-extensively bred on a forest-shrub-prairie habitat divided in different plots separated by high-wire fencing. Animals are bred in separate batches according to sex and age. The number of deer in the estate is around 500 hinds and 100 stags with slight inter-annual variations. Deer are kept within large fenced (6-8 ha) enclosures in batches of 60-80 females; males are kept in separate enclosures. Individual deer identification is performed by ear tagging at weaning.

Calves are managed for weaning at 3.5 months of life and afterwards for sanitary reasons at 6-7 months of age (November-December). Yearlings are managed for sanitary reasons when they are 13 months old (June). At 16 months of age, yearlings are introduced to adult batches and follow the management calendar scheduled for adult deer. Adult deer are managed several times a year, once in winter (January-February) and two-to-three times in summer (July, August and September), for sanitary or productive reasons.

Management schemes of the farm are designed to avoid inducing excessive stress to animals, especially during critical stages of their productive cycle such as calving. The sanitary status of animals in the farm is monitored along the year according to farm health schemes and to existing Spanish and EU laws on notifiable disease surveillance.

234

Table 1. Sample size through sample type and time according to the allocation of animals to vaccinated and unvaccinated groups.

History of C. burnetii infection in the farm

The farm has been found seropositive to C. burnetii in consecutive studies (Ruiz-Fons et al., 2008; González-Barrio et al., 2015 TED, AEM). Reproductive failure occurs in the

235 farm annually, but its rate fluctuates between years (the authors, non-published data). In the 2011 calving season, 27 of 350 (7.7%) pregnant hinds that were confirmed by echography in the last third of gestation failed to breed a calf. This constituted an outstanding rate of reproductive failure in the farm and it is also a high rate of reproductive failure when compared to other studies (Woodbury et al., 2006). In September 2011, around 5 months after the calving season, blood samples and vaginal swabs were collected from pregnant confirmed hinds that failed to breed a calf and from hinds that calved normally (González-Barrio et al., 2015a). Coxiella burnetii was detected in vaginal secretions five months after the calving season. Prevalence in vaginal secretions was higher in females experiencing calf loss (40%) than in females that bred normally (15.4%) and the result was similar for the presence of specific C. burnetii antibodies analyzed by

ELISA (50% and 23.1%, respectively). On average, 17 out of 86 vaginal swabs analysed

(19.7%; 95% CI: 11.9-29.7) were positive to the presence of C. burnetii DNA. The causal effect of C. burnetii infection on reproductive failure could not be confirmed because deer hinds give birth to calves in forest-shrub protected areas within farm plots and are not disturbed until calves are 2-3 months old. Therefore, access to parturition/abortion samples is not possible because disturbance of hinds at this stage would cause a high percentage of calf losses due to abandonment by their stressed mothers.

Design and implementation of the experimental vaccination trial

The endemic status of C. burnetii in the deer farm constituted a unique opportunity to design an experimental program of vaccination in field conditions and evaluate its effectiveness. The experimental trial was designed with the objective of reducing the burden of C. burnetii shed by infected animals and, therefore, reducing infection pressure and avoid potential reproductive effects on deer. For the design of the vaccination program we took into consideration that its implementation should be linked to the usual

236 management schedule of animals in the farm to avoid over-management. Excessive management could be counterproductive because farmed deer still keep most of their wild behaviour and get stressed during management.

An inactivated phase I vaccine that has been widely assayed in European domestic ruminants (Coxevac, CEVA Santè Animale, France) was selected for the field vaccination experiment. Coxevac is commercially available in Spain and its use in mammal species is approved by the European Medicines Agency. The manufacturer recommends a 4ml dose of Coxevac for cattle, 2ml for goats and 1ml for sheep. No previous vaccination trial existed in red deer with Coxevac and therefore we selected the dose we estimated more appropriate according to deer average weight. An intermediate dose (3ml) was selected for deer females because they’re heavier than goats but lighter than cattle. The weight of adult red deer females in the study farm ranges 120-160kg. The vaccine was injected subcutaneously with an automatic injector (Serena 5TPFS, Pimex,

Vizcaya, Spain). We programmed the first vaccination of yearling reposition females to coincide with the sanitary management that is scheduled when they are 13 months old.

According to suggestions of the manufacturer, both adult and yearling females were revaccinated 3 weeks after being given the first dose (Astobiza et al., 2013). Previous experimental vaccination studies in domestic ruminants suggest that animals should be revaccinated every 9-12 months (Astobiza et al., 2013; Rodolakis et al., 2009). We preliminary designed a protocol with annual revaccination coinciding with existing management schemes in the farm. However, in 2013 we analysed long-time series data on the serological status of C. burnetii in the farm and realized that the average half-life of antibodies after natural exposure was around 5-6 months. This finding suggested that any protection linked to humoral immunity would require from revaccination every 6 months (see González-Barrio et al., 2015 FVS). Therefore, we slightly modified the

237 vaccination protocol in 2014 to promote biannual revaccination. The first dose of vaccine was given to adult females (i.e. borne in 2010 or before and consequently named ≤2010 cohort) by mid-January 2012. Most females in this cohort were pregnant when vaccinated. Three-hundred and twenty of the 441 females that were in the cohort ≤2010

(72.6%) were vaccinated and the rest (n=121, 27.4%) were left as control. We allocated vaccinated and control animals to each existing batch in the farm to evaluate the effect of vaccination in coexisting non-vaccinated mates. Therefore any female batch in the farm contained both vaccinated and non-vaccinated animals. Vaccinated animals were revaccinated 3 weeks later at the beginning of February 2012. This cohort of animals was revaccinated 12 months after the first dose of vaccine (January 2013) and thereafter animals were revaccinated biannually until January 2014. The cohort of reposition females borne in 2011 was vaccinated (93 of 124 animals, 75.0% of the 2011 cohort) in

June 2012 for the first time and revaccinated 3 weeks later (Fig. 1). Thirty-one animals

(25.0%) were left unvaccinated and constituted the control group in this deer cohort.

Vaccinated deer were revaccinated 12 months later (July 2013) and subsequently revaccinated biannually. Finally, the cohort of reposition females borne in 2012 was vaccinated (104 of 134 animals, 77.6%) in June 2013 and revaccinated 3 weeks later.

Thirty of the females in this cohort (22.4%) were left unvaccinated. Revaccination was performed in this cohort biannually until January 2014. A descriptive summary of the vaccination protocol applied to each cohort - ≤2010, 2011 and 2012 – is provided in Table

1 and in Fig. 1.

The experiment was approved by the Research Ethics Commission of Castilla – La

Mancha University Animal Ethics Committee.

Monitoring of vaccination effectiveness

238

Coxevac has shown potential to reduce the prevalence of ruminant females that shed C.

burnetii and also to reduce the burden of C. burnetii shed by infected animals (Arricau-

Bouvery et al., 2005; Astobiza et al., 2013; Hogerwerf et al., 2011). We evaluated the

effectiveness of the vaccination trial by collecting vaginal secretions, milk and faeces at

different times - 2.5 (July), 3.5 (August) and 4.5 (September) months - after calving in

2012, 2013 and 2014 (Table 1, Fig. 1). Samples were randomly collected from a subset

of females from each cohort and from both vaccinated and control groups. The survey

was carried along the scheduled management of reproductive females after calving.

Sterile cotton swabs were used to collect vaginal secretions. Milk was extracted by hand

into sterile tubes after disinfecting deer nipples with chlorhexidine and discarding the first

three milk shots. Faeces were collected directly from the rectum with sterile disposable

latex gloves. Vaginal swabs, milk and faeces were transported refrigerated to the

laboratory and preserved frozen at -20ºC until being analysed.

Figure1. Vaccination and sample collection schedule

239

Additionally, to estimate the status of C. burnetii in both vaccinated and non-vaccinated females before the start of the vaccination program and afterwards, blood was collected from the jugular vein into sterile 10ml tubes without anticoagulant (Table 1, Fig. 1). Blood was transported at 4ºC to the laboratory, centrifuged at 3,000g for 10 min and the serum obtained was preserved at -20ºC until analyses were performed. Blood was also collected to estimate seroconversion rates of vaccinated females. Blood samples were collected from hinds of the ≤2010 cohort at 12 different times between 2010 and 2015, 8 times between 2011 and 2015 from the 2011 cohort and 6 times - between 2012 and 2015 - from the 2012 cohort (Fig. 2).

Molecular analyses

DNA from vaginal swabs, milk and faeces was extracted with a commercial DNA extraction and purification kit (DNeasy Blood & Tissue kit, Qiagen, Germany) following the protocols provided by the manufacturer

(http://mvz.berkeley.edu/egl/inserts/DNeasy_Blood_&_Tissue_Handbook.pdf). DNA extraction from swabs was optimized by keeping swabs at 56ºC for 30 min in a solution containing 20 µl of proteinase K and 200 µl of AL buffer. Swabs were thereafter vortexed vigorously for 15 s, removed from tubes and discarded. The remaining sample remained for 30 additional min at 56ºC. After that, the manufacturer's blood extraction protocol was followed. Each sample of milk (200 μl) was mixed directly with ATL and proteinase K and incubated for 3 h at 56ºC; then, the manufacturer's blood extraction protocol was followed. One gram of each faecal samples were mixed with 4 ml of TE buffer (Tris Base

10 mM, EDTA 1 mM, pH 8), vortexed for 30 s and centrifuged at 3,000 g for 2 min.

Thereafter, 200 μl of the supernatant were treated with proteinase K (20 μl) and ATL.

240

) deer groups along the the along groups ) deer iation from the men for average for the average men from iation

Statistically significant differences (p<0.05) between vaccinated and unvaccinated groups unvaccinated and vaccinated between (p<0.05) significant differences Statistically

* * . .

Evolution of seroprevalence (%) and average antibody levels (SP) in vaccinated (black line) and unvaccinated (grey line (grey unvaccinated and line) (black (SP) vaccinated in levels average antibody (%) and seroprevalence of Evolution

implementation of the vaccination experiment. Error bars represent 95% confidence interval of seroprevalence and standard dev standard and seroprevalence of interval 95% confidence represent Error bars experiment. the vaccination of implementation vaccinated were animals at the dates which depict dots Red levels. antibody Figure 2. Figure 241

The concentration of DNA in each aliquot was quantified (NanoDrop 2000, Thermo

Scientific, Waltham, MA, U.S.A.) and frozen at -20ºC until PCR performance. In order to prevent and detect sample cross-contamination, negative controls (Nuclease free water;

Promega, Madison, WI, U.S.A.) were included every 10 samples during the DNA extraction procedure. DNA samples were analysed by a real time PCR (qPCR) targeting a transposon-like repetitive region of C. burnetii as previously described (Tilburg et al.,

2010, González-Barrio et al., 2015a). SsoAdvanced™ Universal Probes Supermix

(BioRad, USA) was used in qPCR according to the specifications of the manufacturer.

DNA extraction and PCR were performed in separate laboratories under biosafety level

II conditions (BIO II A Cabinet, TELSTAR, Spain) to avoid cross-contamination. We extracted C. burnetii DNA from Coxevac and used this as qPCR positive control. Samples were considered positive to the presence of C. burnetii DNA at a threshold cycle (Ct) below 40.0 (Tilburg et al., 2010).

Serological analyses

The presence of specific antibodies against C. burnetii phase I and II antigens in deer sera was determined by using a commercial indirect ELISA test (LSIVet™ Ruminant Q Fever

Serum/Milk ELISA Kit, Life Technologies, USA) with an in-house modification in the secondary antibody (Protein G−Horseradish peroxidase, Sigma-Aldrich, USA) that was previously validated for wild ungulate sera (González-Barrio et al., 2015a). ELISA results were expressed as the sample-to-positive control ratio (SP). For each sample, the SP was calculated according to the formula:

(ODs - ODnc) SP = x100 (ODpc - ODnc) where ‘ODs’ is the optical density of the sample at a dual wavelength 450-620 nm,

‘ODnc’ is the optical density of the negative control and ‘ODpc’ is the optical density of

242 the positive control. All SP values ≤40 were considered as negative, whereas S/P values

>40 were considered as positive. The SP ratio was considered as a proxy of the level of antibodies against C. burnetii as suggested by the manufacturer.

Statistical analyses

Statistical analyses were carried out to compare prevalence and shedding levels of C. burnetii in vaginal swabs, milk and faeces in vaccinated versus unvaccinated hinds and thus to test our hypothesis that deer immunized with inactivated phase I vaccines would experience a reduction in Coxiella burnetii shedding. The humoral immunological response to vaccination in vaccinated females in comparison to the status of unvaccinated animals was evaluated by comparing both seroprevalence and average antibody levels in both groups of deer.

Chi-square tests were employed to compare prevalence of Coxiella burnetii DNA in vaginal swabs, milk and faeces and seroprevalence between vaccinated and unvaccinated groups at each sampling time. Mann-Whitney U non-parametric tests were run to compare burdens of shed C. burnetii (Average Ct) in vaginal swabs, milk and faeces and antibody levels (Average SP) in serum between vaccinated and unvaccinated groups. Finally, to assess for the effect of time on the excretion of C. burnetii and in the serological evolution of the level of antibodies in vaccinated and unvaccinated groups we performed Spearman correlations.

Statistical analyses were run in IBMS SPSS v22.0 (IBM, Armonk, NY, USA). Ninety- five per cent confidence intervals (95%CI) were estimated for prevalence values according to the expression 95%C.I. = 1.96 [p(1 - p)/n]1/2 (Martin et al., 1987), being ‘p’ the proportion in unitary value and ‘n’ the size of the sample employed to estimate the proportion.

243

Results

Seroconversion after vaccination

Two-hundred and two hinds of the ≤2010 cohort that were vaccinated in January 2012 were negative by ELISA prior to vaccination. Three weeks after vaccination, in February

2012, 187 of these hinds (92.6%) seroconverted. In July 2012, 5 months after revaccination, 170 of these females could be analysed again by ELISA and all (100.0%) were positive to C. burnetii antibodies. In contrast, of the 72 animals of the control group in this cohort that were seronegative in January 2012, 39 (54.2%) had seroconverted in

February 2012 and in July 2012 the 69.7% (23/33) of them had seroconverted due to natural infection. The effect of vaccination on activating the humoral response in deer was also evident by the increase in the average level of antibodies. Average SP for deer of the ≤2010 cohort that were seronegative in January 2012 was 14.4 before vaccination and thereafter it experienced a tenfold increase in February 2012 (132.1) and remained similar (121.9) in July 2012. In contrast, non-vaccinated seronegative deer displayed average SP values of 14.9 in January 2012, 86.5 in February 2012 and 65.7 in July 2012.

The 90.0% of seronegative deer from the 2011 cohort that were vaccinated in June 2012

(9/13) had seroconverted 3 weeks later (July 2012), but this rate diminished to 65.2%

(15/23) when analysed in January 2013. Unvaccinated animals did not experience seroconversion in July 2012 (0/11) and in January 2013 (0/11). The level of antibodies experienced a five-fold increase from June 2012 (21.7) to July 2012 (99.1) in the group of vaccinated deer and it slightly decreased by January 2013 (77.1). In contrast, the pattern in unvaccinated animals remained similar along this period with average SP=19.8 in June

2012, 21.7 in July 2012 and 8.6 in January 2013.

244

Finally, seroconversion in the 2012 cohort couldn’t be estimated because serum samples

from seronegative animals allocated to the vaccinated group were not collected in July

2013 - 3 weeks after vaccination; the 20.0% of these animals (3/15) had antibodies in

January 2014. Among seronegative animals of the 2012 cohort control group, 4 of 12

(33.3%) were seropositive in July 2013 and 6 of 15 (40.0%) had antibodies in January

2014. Average SP in the vaccinated group increased from 12.7 in June 2013 to 34.4 in

January 2014. Average SP values were similar in seronegative individuals of the

unvaccinated group: 9.7 in June 2013, 27.0 in July 2013 and 32.8 in January 2014.

Figure 3. Shedding prevalence in vaginal secretions, milk and faeces in vaccinated and unvaccinated deer groups with time from vaccination according to the studied deer cohorts. * Statistically significant differences (p<0.05) between vaccinated and unvaccinated groups.

245

Evolution of the humoral immunity after vaccination

Five-hundred and ninety-five hinds were surveyed for serum at least once between

December 2010 and January 2015 (Table 1). In total, 2194 serum samples were analysed by ELISA. After vaccination and re-vaccination of the ≤2010 cohort in early 2012, seroprevalence remained most of the time around 100% in contrast to the control group in which seroprevalence was always below 20% (Fig. 2A). Statistically significant differences in prevalence between vaccinated and unvaccinated groups in this cohort were found at different times after initiating the vaccination trial (Fig. 2A). The pattern observed in average antibody levels was similar to that observed for prevalence (Fig. 2B); vaccinated animals displayed average SP values over 200.0, an unusually high value in naturally infected ruminants (Ruiz-Fons et al., 2011 Epidemiol Infect). Differences in antibody levels between vaccinated and unvaccinated hinds after vaccination were always statistically significant. Seroprevalence and average antibody levels displayed statistically significantly time trends in the group of vaccinated animals: rho=0.848, p<0.001 and rho=0.902, p<0.001, respectively. Seroprevalence in unvaccinated deer also displayed statistically significant time trend (rho=0.692, p<0.05) although that trend was not clear when represented in a chart (Fig. 3). There was no statistically significant time trend in average antibody levels in this group.

Differences in seroprevalence and average antibody levels between vaccinated and unvaccinated deer of the 2011 and 2012 cohorts were statistically significant after the start of vaccination except in January 2014 (Fig. 2C, 2D, 2E and 2F). There was a statistically significant time trend in both seroprevalence (rho=0.778, p<0.05) and antibody levels (rho=0.762, p<0.05) in the group of vaccinated animals within the 2011 cohort that was not observed in the group of unvaccinated mates. In contrast, statistically significant time trends in seroprevalence (rho=0.886, p<0.05) and antibody levels

246

(rho=0.943, p<0.05) were found in the group of unvaccinated deer of the 2012 cohort but not in vaccinated ones.

Patterns of Coxiella burnetii shedding after vaccination

A total of 444 vaginal swabs, 272 milk samples and 316 faecal samples from 319 hinds

(228 vaccinated and 91 unvaccinated) were investigated during the years 2012, 2013 and

2014 (Table 1). Shedding patterns of C. burnetii in vaginal secretions, milk and faeces were separately analysed for any of the studied cohorts.

Coxiella burnetii DNA was detected in vaginal swabs, milk and faeces in any of the three time periods from calving in which samples were collected, that is up to close the 5th month after calving (Fig. 3). This pattern was similar for any of the three cohorts studied.

Almost no significant differences in shedding prevalence and in average qPCR Ct values were observed between vaccinated and unvaccinated animals in any cohort (Figures 3 and 4). In general terms, shedding prevalence in vaginal swabs (Fig. 3), although not the burden of shed bacteria (Fig. 4), was slightly higher in August than in July and September in contrast to shedding prevalence in milk and faeces (Fig. 3) that tended to decrease along with time from calving. This pattern was observed both in vaccinated and unvaccinated groups in every cohort. An interesting observation was the absence of vaginal shedding in September 2014 in any of the studied cohorts and in both vaccinated and unvaccinated groups. Shedding prevalence of C. burnetii in faeces, although not shedding burden, displayed a decreasing trend with time in vaccinated (rho=0.883, p<0.001) and unvaccinated (rho=0.870, p<0.001) animals of any of the studied cohorts

(Figs. 3 and 4).

Discussion

The use of inactivated phase I vaccines is widely generalized in domestic ruminant populations as a tool to reduce the excretion of C. burnetii by infected animals, and

247 therefore to reduce the burden of bacteria shed into the environment and consequently the risk of infection of humans and other animal species (Arricau-Bouvery et al. 2005;

Guatteo et al., 2008). Recent studies confirmed the efficacy of vaccination for preventing

C. burnetii shedding in uninfected non-pregnant cattle (Guatteo et al., 2008; Taurel et al.,

2012), in uninfected pregnant goats (Arricau-Bouvery et al., 2006) and infected non- pregnant sheep (Astobiza et al., 2013). For optimal vaccine efficacy in different species of domestic ruminants, vaccinated populations may accomplish one common pre- requisite, C. burnetii shall not be present in the population or shall only circulate at low seroprevalence. Additionally vaccination at early ages, when animals have not yet been exposed to infection by C. burnetii, is recommended.

Current demographic status of red deer populations in the Iberian Peninsula in addition to the widespread geographic distribution of C. burnetii in this species and the increasing relevance of deer as a game and farm species that increases red deer-human interaction rates makes research on potential control measures of C. burnetii transmission a high relevance issue. In our study, we selected a semi-extensive bred red deer population as a model to test the efficiency of vaccinating red deer against C. burnetii infection with a phase I inactivated vaccine (González-Barrio et al., 2014; González-Barrio et al., 2015 submitted Frontiers Vet Sci). Since C. burnetii is endemic in around 50% of red deer populations in Iberia (González-Barrio et al., 2015 AEM), analysing the efficiency of the application of the vaccine to an endemic population would better resemble what we could expect from vaccinating free-roaming deer population. We aimed to simulate in a semi- controlled red deer population what would be the effect of vaccinating a high percentage of deer in a free-roaming population with endemic circulation of C. burnetii. Wildlife vaccination chances are increasing as long as research on efficient oral vaccines and vaccine delivery methods to wildlife progress (Beltrán-Beck et al., 2014; Gortázar et al.,

248

2014). Therefore, research on potential control tools should be promoted. Besides studying the effect of vaccination on the excretion of C. burnetii in red deer, this study

also explores the main shedding routes of C. burnetii in this wild species.

ically ically

Statist

* *

roups.

shed (average qPCR Ct value) and associated standard deviation (error bars) in vaginal secretions, milk and faeces faeces and milk secretions, vaginal in bars) (error deviation standard associated and value) Ct qPCR (average shed Coxiella burnetii burnetii Coxiella

of Burden

significant differences (p<0.05) between vaccinated and unvaccinated groups unvaccinated and vaccinated between (p<0.05) differences significant by by any cohort of deer studied according to their allocation to vaccinated (black diamonds) and unvaccinated (grey diamonds) g 4. Figure

249

Humoral immune response to vaccination

In general terms a high percentage of seronegative red deer seroconverted after vaccination with Coxevac, with apparently higher seroconversion rates in adult than in yearling individuals. This age-related effect on humoral response has been also observed in natural infection patterns by C. burnetii in the study farm (González-Barrio et al., 2015

FVS). This seems to be related to increasing immune competence with individuals’ age.

Vaccination and subsequent revaccination induced a high and stable seroprevalence in the population that remained high when animals were revaccinated both annually and biannually. Vaccination with a single dose (first vaccination) induced high humoral response in red deer in a short period of time (3 weeks). Although the boosting effect on the humoral immune response after revaccination could not be evaluated in the short time, the level of antibodies remained similar few months later. This suggests that perhaps revaccination 3 weeks after first vaccination is not necessary in red deer and that revaccination every 6 months, as previously suggested (González-Barrio et al., 2015

FVS), would keep high levels of circulating antibodies in vaccinated individuals. In domestic ruminants changes in the level of C. burnetii antibodies after vaccination are similar, with 95% seroconversion rates observed in goats 28 days after vaccination

(Arricau-Bouvery et al., 2005). However, in vaccinated goats antibodies last between 8 and 12 months (Arricau-Bouvery et al., 2005; Coxevac data sheet) in contrast to what we observed in vaccinated and revaccinated deer. Seroconversion rates observed in vaccinated sheep are variable among studies: 40%, 98% and 100% (Astobiza et al., 2011;

Eibach et al., 2013; Hamann et al., 2009). Brooks et al. (1986) detected antibodies in sheep vaccinated with a phase I vaccine (not Coxevac) until 11 months after vaccination.

In adult cattle, a low proportion of the animals need annual revaccination while the 80% of them maintained antibodies one year after vaccination (Rodolakis et al., 2009). This

250 pattern varies with cattle age and only in the 68% of vaccinated yearling cows antibodies were detected one year after vaccination. Antibody levels in cattle heifers induced by vaccination also had lower average half-life than these in adult cattle (Rodolakis et al.,

2009).

Data obtained in this study shows that vaccination and revaccination of deer with phase I inactivated vaccines induces long-lasting humoral response in a high percentage of individuals. However, revaccination of adult and yearling females in July 2013 did not induce the maintenance of existing antibody levels and seroprevalence also decreased by

January 2014. This was observed in the three studied cohorts, which suggests it may be related perhaps to the conservation of the vaccine or to a failure of the vaccine batch provided by the manufacturer. However, both explanations have been rejected after checking that vaccines used in July 2013 were from the same batch than those employed in June 2013 in yearlings of the cohort 2012. These animals mounted a normal humoral immune response after vaccination in June 2013. Vaccines were preserved in refrigerated conditions with no recorded change in the temperature of the refrigerator - which is surveyed three times a week - in which these were kept from June to July 2013. We found no plausible cause for that phenomenon.

Effects of vaccination on Coxiella burnetii shedding patterns in red deer

Coxiella burnetii DNA was detected in vaginal swabs of both vaccinated and unvaccinated hinds until 4.5 months after calving. This shedding pattern in vaginal secretions resembles that reported in infected vaccinated and non-vaccinated goats

(Rousset et al., 2009). Vaccination did not reduce the burden of C. burnetii shed in vaginal secretions along the study period but a reduction in the time the pathogen was shed in vaginal secretions was observed in the third year from the beginning of the vaccination

251 program. A longer monitoring period would be perhaps needed to properly evaluate the effect of vaccination over reduction of C. burnetii shedding as previously suggested for endemically vaccinated sheep populations (Astobiza et al., 2011b). In vaccinated uninfected goats Arricau-Bouvery et al. (2005) observed a reduction in vaginal shedding time just two weeks after vaccination. Although not every vaccinated yearling deer in

2011 and 2012 cohorts had been exposed to C. burnetii by the time the vaccination was implemented (at 13 months of age; Fig. 2), we did not observe a general reduction of the shedding time in a short time window. However, since our results come from a field experiment in deer within a highly contaminated environment (see González-Barrio et al., 2015 AEM y EID) and since we observed a reduction in vaginal shedding time with vaccination, we cannot discard that vaccination of uninfected deer under low natural infection pressure conditions would significantly reduce vaginal shedding and in a shorter time period. We did not observe any reduction in the burden of C. burnetii shed in vaginal secretions, but we have to remark that shedding burdens were always at high Ct values and very few animals displayed Ct values in qPCR below 30.0. Low C. burnetii burdens have been reported in naturally infected red deer before (González-Barrio et al., 2014

AEM). These findings contrast with what it has been observed in goats after vaccination

(Hogerwerf et al., 2011). In naturally infected sheep populations with ongoing vaccination along 3-4 years, the percentage of shedders and shedding burdens in vaginal secretions decreased with time and even disappeared (Astobiza et al., 2013; Astobiza et al., 2011). However, there are methodological differences among studies that make comparisons difficult.

Coxiella burnetii shedding in milk lasted a little bit less than vaginal shedding in infected deer. This finding was consistent along the studied deer cohorts in which shedding was approximately restricted to the first three months from calving. No differences in this

252 pattern were evidenced between vaccinated and control groups. Milk shedding in unvaccinated naturally infected goats has been observed until 4-6 weeks (Arricau-

Bouvery et al., 2005; Roest et al., 2012). Vaccination of goats reduced the bacterial load and time of C. burnetii in milk (Hogerwerf et al., 2011). The shorter time of milk shedding compared to vaginal secetions or faeces agrees with previous reports in sheep and goats

(Rodolakis et al., 2007; Astobiza et al., 2010; Roest et al., 2012). The observed absence of reduction in shedding prevalence and burden of C. burnetii in vaccinated deer agrees with what it has been observed in sheep and goats in some experimental vaccination studies (Rodolakis et al., 2007; Astobiza et al., 2010; Roest et al., 2012; Astobiza et al.,

2013).

Coxiella burnetii shedding in faeces was detected 4.5 months after calving as previously observed in non-vaccinated goats (Roest et al., 2012). The main finding of our study was the progressive reduction of C. burnetii shedding prevalence in faeces, although not the burden of shed bacteria, with time from the implementation of vaccination. This was observed both in vaccinated and unvaccinated deer groups but, since animals in both groups were mixed in existing batches, the reduction observed in unvaccinated animals may be a consequence of the reduction of the burden of C. burnetii shed in faeces by vaccinated deer. A reduction in environmental contamination caused by the reduction of shedding prevalence in faeces - if we assume faeces may be the main source for environmental contamination and transmission of C. burnetii as previously suggested in cattle (Courcoul et al., 2011) - would have accounted in a global reduced infection pressure in the farm that would have been observed by a reduction in the prevalence of exposed animals within the control group. However, this was not observed in the studied cohorts. A recent study on the dynamics of C. burnetii in the deer of the study population found a decreasing trend of incidence in yearling females from 2012 to 2014 along the

253 implementation of the vaccination program (González-Barrio et al., 2015 FVS). If the observed decrease in C. burnetii shedding prevalence in faeces is caused by vaccination and if it is related to decreasing infectious pressure in the farm cannot be proved with the data we gathered in this study. Therefore, further studies should evaluate in the long-time these potential effects of vaccination in deer.

Conclusions and recommendations

Vaccination trials in endemic sheep populations concluded that longer vaccination periods are needed in ruminants to observe any effect of the vaccination in reducing C. burnetii shedding prevalence and burden, and therefore to reduce infection pressure within the population (Astobiza et al., 2011). We initiated the experimental vaccination trial in a deer population in which C. burnetii was endemic and where a high percentage of the animals had been exposed to infection by C. burnetii (González-Barrio et al., 2015

FVS). However, since we aimed to test the efficiency of implementing vaccination in naturally infected endemic populations to resemble what we could find in the real world when planning measures to reduce the risk of transmission from free-roaming red deer populations to livestock and/or humans, we decided to carry out the field experiment in a model deer population. Unfortunately, the costs of implementing a vaccination experiment over such a big population and monitoring the effect on shedding are extremely high and therefore long-time monitoring is difficult to achieve at such scales.

We faced potential drawbacks to the effect of vaccination such as the endemic status of

C. burnetii and the initial vaccination of pregnant animals (Guatteo et al., 2008; Rousset et el al., 2009, de Cremoux et al., 2012). In pregnant dairy cattle, Guatteo et al., (2008) observed that the likelihood of shedding was similar in vaccinated that in control animals.

Future vaccination trials may perhaps target non-pregnant deer to avoid any potential effect of pregnancy on the immune response of hosts to vaccination. Start the vaccination

254 trial targeting only young animals while leaving adult animals unvaccinated could be an approach to evaluate in the future. However, in our opinion, that approach would need from an increased time of vaccination to achieve any potential reduction in C. burnetii shedding in the population. We herein decided to start vaccinating yearling animals at 13 months of age for the first time according to preliminary observations on natural exposure to C. burnetii with age in the study population. However, after epidemiological analyses were completed in the study farm for a long time period (González-Barrio et al., 2015

FVS), we would recommend that future vaccination programs target calves at 6-7 months of age to receive the first vaccine dose. That would perhaps protect them in the time window between the loss of maternally-derived antibodies (González-Barrio et al., 2015

FVS) and the main shedding period in deer populations when they around 12-13 months old.

Acknowledgements

We thank deer farm keepers for their valuable help in deer management and sample collection. This work was funded by EU FP7 Grant ANTIGONE (278976) and CDTI

(Centro para el Desarrollo Tecnológico Industrial, Spanish Ministry for the Economy and

Competitiveness - MINECO). FRF is supported by a ‘Ramón y Cajal’ contract from

MINECO.

255

Capítulo V. Síntesis y Conclusiones

256

Síntesis

Este capítulo resume los resultados más relevantes de los diferentes trabajos de investigación que componen la presente Tesis Doctoral sobre la epidemiología y el control de Coxiella burnetii en la fauna silvestre, haciendo especial énfasis en el impacto que los conocimientos adquiridos en los trabajos realizados pueden tener sobre el sector cinegético, sobre el sector ganadero y sobre el diseño de futuras estrategias de prevención y control de C. burnetii en la fauna silvestre por parte de las autoridades en materia de conservación de la biodiversidad y la fauna silvestre, la sanidad animal y la salud pública.

Se mostrará el potencial como reservorio de C. burnetii de las especies silvestres objeto de estudio y las implicaciones de los hallazgos, se analizarán los resultados sobre la dinámica de C. burnetii en escenarios endémicos en sistemas de producción de ciervo rojo en extensivo y cómo estos resultados pueden ser aplicados al control de la infección, se analizará la aplicabilidad de los avances en el conocimiento sobre la epidemiología molecular de este patógeno sobre el conocimiento de su dinámica y de las relaciones con los hospedadores a los que infecta, se analizarán los resultados que han conllevado a estimar las vías de excreción y transmisión de C. burnetii en las especies silvestres objeto de estudio y su aplicabilidad para profundizar en el conocimiento de la epidemiología del patógeno y el desarrollo de medidas de prevención y control, y se propondrán medidas de control de la infección por C. burnetii en ciervo rojo basadas en los resultados obtenidos tras el diseño, aplicación y evaluación de un protocolo de vacunación con vacunas comerciales inactivadas.

El incremento durante las últimas décadas de la concienciación en la sociedad por la conservación de la naturaleza, por seguir hábitos alimentarios más saludables y sostenibles, por la práctica de deportes relacionados con la naturaleza, así como el

257 aumento en la ganadería y el incremento por el interés en actividades cinegéticas, entre muchos otros motivos, han propiciado una mayor frecuencia de contacto entre las especies silvestres, el ganado y el ser humano que puede generar conflictos y favorecer la transmisión de enfermedades infecciosas compartidas con animales silvestres. Conocer cuál es el papel de las especies de fauna silvestre con las que el ser humano puede tener una mayor frecuencia de contacto - especies con amplia distribución geográfica, abundantes y con interés cinegético - en el mantenimiento y la transmisión de C. burnetii, identificar los factores que condicionan el mantenimiento de C. burnetii en las poblaciones de fauna silvestre, conocer las relaciones patógeno-hospedador que determinan la dinámica del patógeno y su transmisión, así como evaluar potenciales medidas de control de la infección en especies silvestres son aspectos esenciales para prevenir la transmisión de C. burnetii desde la fauna silvestre a los animales domésticos y al ser humano. Sólo los conocimientos científicos adecuados y el desarrollo y ensayo de estrategias de control pueden prepar a las sociedades humanas para evitar los riesgos sanitarios que su contacto con la fauna silvestre conllevan.

Coxiella burnetii circula de forma endémica y a gran escala en especies silvestres

ampliamente distribuidas en la península ibérica

como el ciervo rojo y el conejo de monte

Numerosas especies de animales silvestres presentan una distribución geográfica amplia en la península ibérica, pero de todas ellas aquellas con interés cinegético pueden representar un riesgo mayor en la transmisión de C. burnetii a los animales domésticos y al ser humano por la esperable mayor tasa de interacción. Especies cinegéticas como el ciervo rojo, el conejo de monte o el jabalí representan las mayores presas cinegéticas en la España peninsular de caza mayor y menor, respectivamente. Sus poblaciones presentan una distribución geográfica amplia, ocupando casi toda la península ibérica en su

258 conjunto, son apreciadas como trofeo cinegético, se consume su carne, muchas de sus poblaciones están sometidas a algún grado de gestión cinegética y coexisten con ganado doméstico. Si alguna especie silvestre puede representar algún riesgo para el ganado y el ser humano como reservorio de C. burnetii, estas especies deben, a priori, estar entre ellas.

Así, tres de los estudios de esta tesis se enfocaron a estudiar la presencia de C. burnetii en estas especies, si bien en el caso del jabalí su estudio se limitó a poblaciones densas del centro-sur peninsular en las que las prevalencias de circulación de otros patógenos son altas. Se obtuvo una información representativa del estado de C. burnetii en las poblaciones de ciervo rojo (CAPÍTULO II.1) y conejo de monte (CAPÍTULO II.2) en la península ibérica, incluyendo también poblaciones de Portugal. El 50% de las poblaciones de ciervo estudiadas y más del 60% de las poblaciones de conejo presentaron exposición a C. burnetii. Además, se detectó la circulación de C. burnetii en poblaciones de estas especies en granja; el 67% de las granjas de ciervo y el 50% de las de conejo tenían presencia de C. burnetii. Al contrario de lo esperable debido a las altas tasas de prevalencia de otros patógenos que el jabalí comparte con animales domésticos, pero coincidiendo con los resultados obtenidos en ciervo para la misma zona, la prevalencia de C. burnetii en los jabalíes analizados de la zona centro-sur de España fueron bajas. Se identificaron algunos factores de riesgo de exposición del ciervo y del conejo a C. burnetii, pero los modelos estadísticos mostraron que otros factores no contemplados en estos estudios deben de tener mayor peso en la dinámica de C. burnetii que los contemplados. Curiosamente, la gestión cinegética del ciervo, al contrario que ocurre con enfermedades como la tuberculosis bovina, no representa un factor de riesgo de exposición del ciervo a C. burnetii. Las poblaciones no gestionadas presentaron tasas más altas de exposición a la bacteria que aquellas gestionadas con fines cinegéticos, si bien las máximas prevalencias se observaron en una granja de ciervos. En una situación

259 especial en la que coexisten conejo y ciervo (en la provincia de Cádiz), la presencia de ciervo mostró una influencia positiva sobre la exposición de los conejos a C. burnetii. A pesar de que la reproducción del conejo se produce durante todo el año en esta zona templada de España, se observó un mayor riesgo en verano que podría estar asociado a un efecto de la presión de infección ejercida por la secreción masiva de C. burnetii tras el parto de ciervas en simpatría (CAPÍTULOS II.3 Y IV). Este resultado sugeriría un vínculo epidemiológico entre ciervo y conejo cuando están en simpatría, sugiriendo que ambas especies comparten las mismas cepas de C. burnetii circulantes en el medio.

Estudios previos realizados a nivel mundial utilizando la fauna silvestre como modelo evidencian que C. burnetii está ampliamente distribuida en todos los ecosistemas del planeta. La presencia de C. burnetii ha sido probada en decenas de animales, desde mamíferos terrestres y marinos, aves, reptiles, anfibios, e incluso artrópodos. Esta característica multi-hospedador confiere a C. burnetii un potencial zoonósico importante que merece ser estudiado en profundidad. Ciervo rojo y conejo de monte, probablemente jabalí, pueden ser especies importantes en el ciclo de vida de C. burnetii en la península ibérica y constituir importantes reservorios para animales domésticos y para el ser humano. Este conocimiento hasta el momento inexistente demuestra la necesidad de considerar la potencial interferencia de la fauna silvestre en cualquier sistema de control que se implemente en el ganado para reducir la prevalencia de C. burnetii, así como considerar a estas especies silvestres como potenciales fuentes de transmisión a humanos y, por ello, responsables de casos clínicos de esta infección. Estos conocimientos son de aplicabilidad, por lo tanto, tanto para las autoridades en materia de sanidad animal como de salud pública en el desarrollo de planes de prevención y control de la fiebre Q en la península ibérica.

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El ciervo rojo y el conejo de monte son reservorios de Coxiella burnetii en la

península ibérica

Tres aspectos pueden hacer del ciervo rojo y del conejo de monte reservorios verdaderos e importantes de C. burnetii: i) Amplia distribución geográfica y capacidad de alcanzar altas densidades poblacionales; ii) Alta prevalencia poblacional de C. burnetii a lo largo y ancho de sus áreas de distribución; y iii) Capacidad de ser infectados, replicar y excretar el patógeno al medio ambiente después de haber sufrido una infección sistémica. Los tres requisitos son cumplidos por ciervo rojo (CAPÍTULOS II.1, III.1 y IV) y conejo de monte (CAPÍTULO II.2) en la península ibérica. Sólo uno de esos aspectos no ha podido ser demostrado en el jabalí (CAPÍTULO III.2) porque no se realizó un estudio a escala geográfica adecuada para estimar la prevalencia de C. burnetii en sus poblaciones, pero la amplia distribución geográfica y las altas densidades poblacionales de las poblaciones ibéricas de jabalíes son ampliamente conocidas y en este estudio (CAPÍTULO III.2) se ha confirmado la capacidad del jabalí para excretar C. burneti.

Los resultados de los estudios serológicos que confirmaron la amplia distribución de C. burnetii en ciervo rojo y conejo de monte en la península ibérica fueron completados con estudios de prevalencia de infección sistémica (detección de ADN de C. burnetii en muestras de bazo; CAPÍTULOS II.1 y II.2) y con estudios sobre las vías de excreción de C. burnetii por parte de estos animales (detección molecular de la presencia de C. burnetii en hisopos genitales, orales, nasales y rectales y en muestras de glándula mamaria/leche y heces; CAPÍTULOS II.2, III.1 III.2 y IV). El 50% de las poblaciones de ciervo rojo, al igual que se observó con el análisis serológico, mostraron presencia de infección sistémica por C. burnetii (muestras de bazo positivas en PCR). Las muestras de bazo analizadas en conejo también señalaron la presencia de infecciones sistémicas por

C. burnetii. Se detectó excrecion de C. burnetii por numerosas de las vías estudiadas en

261 las tres especies, confirmándose en el jabalí la capacidad de los machos de transmitir C. burnetii a través del semen, como recientemente se ha confirmado en moruecos de raza manchega y en gacelas en España.

Estos resultados, junto a los resultados obtenidos en esta Tesis Doctoral sobre la dinámica de infección por C. burnetii en el ciervo (CAPÍTULO II.3), señalan que es posible que el sesgo de los muestreos de ciervo y jabalí a la época principal de caza - otoño e invierno

- puedan conllevar a que las prevalencias de C. burnetii hayan sido subestimadas, siendo en ese caso incluso superiores. Por lo tanto, deberíamos considerar a ambas especies, y potencialmente a especies como el jabalí, como reservorios de C. burnetii en la península ibérica. Estos resultados deben ayudar a mejorar los sistemas de prevención de transmisión de C. burnetii en la interfaz silvestre-doméstico y silvestre-humano, reduciendo así los riesgos.

En poblaciones endémicas de ciervo rojo la infección por Coxiella burnetii es

dinámica y determina variación en la presión de infección en el tiempo

La dinámica a largo plazo de la infección por C. burnetii en rumiantes silvestres y, sobre todo, en rumiantes domésticos no había sido determinada en ningún estudio anterior. En poblaciones de hospedadores en las que un patógeno circula de forma endémica podría establecerse una relación entre la inmunidad de la población (conocida en animales domésticos como inmunidad de rebaño) y la replicación y transmisión del patógeno. Si existe algún efecto de la inmunidad de la población sobre la dinámica del patógeno, éste determinaría variabilidad temporal en la presión de infección por C. burnetii y, por lo tanto, variación en la incidencia de primoinfecciones en individuos susceptibles; consecuentemente, se observaría variación en el tiempo en los niveles de exposición de toda la población. Coxiella burnetii infecta a las hembras de rumiantes silvestres a edades

262 tempranas y ocasiona problemas reproductivos en un porcentaje no muy elevado de las hembras jóvenes del rebaño, y en menor medida en las hembras adultas. Estos hechos sugieren que las hembras de rumiantes adquieren algún tipo de inmunidad tras sucesivas infecciones que protege de los efectos clínicos de C. burnetii y, quizás, reduce la capacidad de replicación del patógeno y su excreción. Cuando en la población se alcanza un porcentaje elevado de animales ‘protegidos’ frente a la infección, la excreción podría verse reducida y, con ello, la contaminación ambiental y la presión de infección. Tras un tiempo y debido a la desaparición paulatina de animales ‘inmunizados’, el porcentaje de animales susceptibles aumentaría y, paralelamente, la excreción, el nivel de contaminación ambiental y, por ende, la presión de infección. Para probar esta teoría son necesarios diversos estudios científicos experimentales tanto en condiciones de laboratorio como en condiciones de campo.

En esta Tesis Doctoral se abordó estudiar si esta teoría podría ser factible utilizando como modelo una población endémica de ciervo rojo (CAPÍTULO II.3) monitorizada en el tiempo. Se estudió la dinámica de exposición a C. burnetii en ciervas de diferentes edades mediante análisis serológico a lo largo de 12 años consecutivos y se analizó el efecto que el porcentaje de animales en la población con inmunidad humoral frente a C. burnetii podría tener sobre la presión de infección experimentada por hembras susceptibles jóvenes en el rebaño. Los resultados obtenidos muestran una situación dinámica en el tiempo de la infección por C. burnetii, con niveles variables de seroprevalencia entre años tanto en ciervas adultas como en ciervas jóvenes. La incidencia en ciervas jóvenes varió notablemente en el tiempo, indicando variación inter-anual en la presión de infección en el rebaño. Los análisis de correlación entre el estado de la inmunidad humoral en la población de hembras adultas y la incidencia en ciervas jóvenes mostraron una relación negativa aunque no estadísticamente significativa. Esto indicaría que a mayores niveles

263 de seroprevalencia en las hembras adultas en la población, menor sería el riesgo de infección para hembras jóvenes, es decir, menor sería la presión de infección. Mientras, en años con menor seroprevalencia en hembras adultas, la presión de infección para las hembras jóvenes sería mayor. Desafortunadamente la escala temporal del estudio parece no ser suficiente para probar la existencia de relación y en el estudio no se pudo analizar la variación en la contaminación ambiental por C. burnetii.

Estos resultados muestran que poblaciones de rumiantes endémicas para C. burnetii, en las que la vacunación con vacunas de fase I inactivadas - la estrategia más efectiva en rumiantes domésticos - no es recomendable, podrían presentar ventanas temporales en las que los niveles de inmunidad humoral global fuesen más bajos y por lo tanto se pudiese acceder así a proteger un mayor porcentaje de animales susceptibles en la población mediante la implantación de la vacunación. Poder predecir cuando ocurren esas ventanas sería muy favorable para el control de la infección en poblaciones endémicas, las cuales constituyen aproximadamente el 50% de las poblaciones de rumiantes en España. Esas ventanas temporales vendrían después de periodos continuados - aunque aún no se conoce su duración - de altos niveles de inmunidad humoral en el rebaño y baja incidencia de primoinfecciones en animales jóvenes. Los resultados obtenidos en este estudio proporcionan una base empírica sobre la que profundizar posteriormente y que podría contribuir al diseño de estrategias más eficaces de control de la infección por C. burnetii tanto en rumiantes domésticos como silvestres.

Las hembras de ciervo rojo son infectadas por C. burnetii al año de vida

Un dato importante que proporcionaría una valiosa información para la implantación de vacunación como método de control de C. burnetii en las poblaciones de ciervo rojo sería conocer cuándo se infectan los animales por primera vez a lo largo de su vida. Así, se

264 podría determinar la edad a la que es recomendable iniciar la vacunación como medida de protección frente a la infección por C. burnetii.

Para determinar este dato, se estudiaron tres cohortes de ciervas - nacidas en los años

2008, 2009 y 2010 - en una población endémica (CAPÍTULO II.3) de forma consecutiva entre los 7 y los 78 meses de vida (algo más de 6 años). Se analizó el estado inmunológico humoral de estas cohortes hasta en 13 ocasiones consecutivas entre los 7 y los 78 meses de vida. La mayoría de los animales eran seronegativos a los 7 meses de vida, pero ese porcentaje aumentaba exponencialmente a partir de los 13 meses de vida, indicando que la infección se produce principalmente a partir del primer año de vida tras la época de partos de las ciervas gestantes en la población.

Este resultado señala que sería recomendable iniciar la vacunación de las ciervas entre los 7 y los 13 meses de vida, antes de la principal época de partos de las ciervas gestantes.

A pesar de que pueden existir diferencias en la dinámica de C. burnetii en una población de ciervo en condiciones ‘controladas’ como la estudiada y una población de ciervos

‘libre’, la población estudiada sigue unos ritmos de vida bastante naturales, con partos naturales no asistidos y monta natural como complemento de la reproducción asistida.

Quizás sería esperable, de acuerdo con las diferencias observadas en el nivel de seroprevalencia entre esta población y las poblaciones ibéricas de ciervo en condiciones de libertad (CAPÍTULO II.1), que incluso el porcentaje de primoinfecciones al año de vida fuese menor en las poblaciones en libertad. Por ello, vacunar a los animales antes de alcanzar su primer año de vida también sería recomendable en estas poblaciones en libertad. Estos resultados serán de utilidad para el diseño de estrategias futuras de control de C. burnetii en ciervos, tanto en condiciones controladas como en libertad. El número creciente mundial de granjas de ciervo señala que estos resultados serán de aplicabilidad para la mejora de las condiciones sanitarias de dichas granjas.

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Altas dosis de anticuerpos podrían proteger frente a la infección por Coxiella

burnetii en ciervo rojo

Hoy en día aún se desconoce qué tipo de inmunidad juega algún papel en la protección frente a C. burnetii en animales. Los resultados de los modelos experimentales en roedores de laboratorio no clarifican si la inmunidad humoral, la inmunidad celular o ambas a la vez son responsables de la protección frente a C. burnetii. Analizar con modelos experimentales controlados cuáles son los mecanismos inmunológicos que protegen frente a la infección por C. burnetii en grandes animales es complicado y costoso. Sin embargo, buscar evidencias epidemiológicas que indiquen la relación entre el estado inmunológico de los animales y la infección podría ayudar a proponer hipótesis que faciliten el diseño de experimentos controlados en este tipo de animales.

En uno de los trabajos llevados a cabo en esta Tesis Doctoral (CAPÍTULO II.3) se realizó un seguimiento longitudinal en una cohorte de animales nacidos en 2013 en una población de ciervo donde C. burnetii es endémica desde su 2º mes de vida hasta casi los

2 años de edad. El seguimiento consistió en el análisis serológico por ELISA de muestras de suero recolectadas a los 2, 3, 7, 13, 14, 19 y 20 meses de edad. Los resultados mostraron elevados niveles de anticuerpos a los 2 meses de edad con un elevado porcentaje de seropositividad en los animales a esta edad. Los anticuerpos habían desaparecido hacia los 7 meses de edad y volvieron a aparecer a los 14 meses, probablemente como consecuencia de infección natural por C. burnetii. Alrededor de los 20 meses de edad tanto la seroprevalencia como el nivel de anticuerpos había disminuido notablemente. Los elevados niveles de anticuerpos observados en las ciervas a los 2 meses de edad comparados con los valores medios a los 14 meses (tras infección natural) sugieren que las ciervas transmiten a sus crías una alta dosis de anticuerpos maternales en la lactación.

Alrededor de los 7 meses de vida estos anticuerpos han desaparecido y sólo vuelven a

266 aparecer alrededor del año de vida tras la principal época de partos de las ciervas gestantes en la población. Aparentemente estos niveles elevados de anticuerpos podrían conferir algún tipo de protección temporal de corta duración frente a la infección por C. burnetii ya que la mayor parte de los animales en el rebaño seroconvierte a partir del primer año de vida, sugiriendo primoinfección a esa edad y no antes. Estos resultados, de ser confirmados con estudios experimentales con mayor control, señalarían que la vacunación de los animales debería de producirse entre los 5 y 7 meses de vida del animal, aunque - como confirman los resultados del estudio de las cohortes 2008-2010 en esta población - siempre antes de la llegada de la época de partos alrededor de su primer año de vida.

Ante la falta de estudios experimentales controlados que proporcionen información de cómo diseñar un protocolo de vacunación en ciervo, estos resultados son de utilidad para diseñar experimentos de vacunación en campo que inmunicen a los animales a edades tempranas justo entre la pérdida de anticuerpos maternales y la primoinfección por bacterias ambientales.

Diferencias en los genotipos de C. burnetii circulantes en ciervo rojo y conejo de

monte en simpatría sugieren algún tipo de adaptación

del patógeno a su hospedador

La información existente sobre los genotipos de C. burnetii que circulan en las especies de fauna silvestre es escasa a nivel mundial y, por ende, conocer si estos genotipos son compartidos entre diferentes especies de hospedadores es difícil. Toda caracterización molecular de los genotipos circulantes en la fauna silvestre sería de utilidad para trazar el origen de brotes en ganado y en humanos con potencial origen en la fauna silvestre.

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Por esta razón, en esta Tesis Doctoral se abordó el genotipado de muestras de diferentes hospedadores silvestres ibéricos que fueron positivas en PCR para C. burnetii

(CAPÍTULO II.4). Se seleccionó un método de genotipado - MLVA - anteriormente usado para tipar un gran número de cepas del ganado doméstico, de casos clínicos de fiebre Q en humanos y de algunos animales silvestres, con la finalidad de que los resultados fuesen comparables. Se logró genotipar de forma completa o casi completa un total de 22 genotipos diferentes presentes en ciervo rojo y conejo de monte. La mayor parte de las muestras genotipadas - todas salvo una - eran originarias de una zona del sur de España donde ciervo y conejo coexisten y comparten hábitat y alimento. Sin embargo, a pesar de este hecho y de que las muestras tipadas en ambas especies fueron recolectadas en los mismos años, se observó una clara separación de los genotipos circulantes en ciervo y aquellos que infectaron al conejo. Los genotipos de ciervo se agruparon aparte de los genotipos de conejo que también presentaron agrupación. Algunos de los genotipos presentes en ciervo y conejo presentaban patrones de MLVA idénticos o muy similares a genotipos aislados en ganado y en casos humanos de fiebre Q, sugiriendo que la fauna silvestre comparte genotipos de C. burnetii con ganado y humanos. Sin embargo, la sorprendente separación de los genotipos en función del hospedador a pesar de que los hospedadores conviven en simpatría, sugiere algún tipo de adaptación de los genotipos de C. burnetii a particularidades del hospedador.

La aplicabilidad de estos resultados es limitada debido a la limitación en el número de muestras que pudieron ser tipadas, al limitado número de especies silvestres en las que se pudo genotipar C. burnetii y al limitado origen geográfico de los genotipos identificados.

Sin embargo, cierta especificidad de hospedador también ha sido sugerida en estudios moleculares en diferentes especies de rumiantes domésticos en simpatría. Este estudio señala la necesidad de profundizar en la caracterización molecular de C. burnetii en

268 diferentes especies domésticas y silvestres y en humanos con una aproximación metodológica mucho más exhaustiva, que abarque un ámbito geográfico mayor y, quizás, integrando diferentes técnicas de tipado molecular. Investigaciones futuras con esta aproximación podrían clarificar si existe algún tipo de adaptación de C. burnetii a sus hospedadores y estimar en qué forma este hecho determina la dinámica de infección por

C. burnetii y qué factores determinan que los genotipos puedan ser compartidos por unas y otras especies. Quizás estos resultados mejoren las capacidades de control de C. burnetii en el futuro.

Las cepas de Coxiella burnetii que infectan a la fauna silvestre son genéticamente

similares a las aisladas en casos clínicos humanos en España

A nivel nacional, en España, el método empleado para el genotipado de C. burnetii en muestras de diferentes hospedadores ha sido la PCR-RLB. Tipar con este método cepas de C. burnetii presentes en la fauna silvestre española sería de gran utilidad para estudios comparativos con la información existente en nuestro país.

Por ello, tras el tipado molecular basado en MLVA (CAPÍTULO II.4) que pretendía una comparación de cepas a la escala internacional a la que la información estaba disponible, se planteó el tipado molecular de las cepas de C. burnetii en la fauna silvestre española mediante PCR-RLB para un estudio comparativo a nivel nacional (CAPÍTULO II.5).

Los resultados mostraron que algunos genotipos aislados en fauna silvestre también han sido descritos en animales domésticos y humanos. Mientras que los genotipos procedentes de rumiantes domésticos en este estudio se agruparon con genotipos anteriormente descritos en ganado en España, los genotipos de fauna se agruparon entre sí y con genotipos de garrapatas y casos clínicos humanos. Estos resultados, conforme a lo que sugieren los resultados de análisis mediante MLVA, sugieren algún tipo de

269 adaptación de las cepas de C. burnetii a sus hospedadores y/o vectores. Aunque los resultados moleculares de las cepas de C. burnetii circulantes en España aún son escasos para sacar conclusiones firmes, el que los genotipos predominantes en fauna silvestre sean también predominantes en garrapatas y en casos de hepatitis aguda humana en España sugieren que quizás las cepas circulantes en un ciclo fauna silvestre-garrapata pueden ocasionalmente ser transmitidas a humanos. Si este ciclo silvestre está detrás de la separación geográfica en la presentación clínica aguda de la fiebre Q en humanos en

España - con cuadro neumónico predominante en el tercio norte y cuadro de hepatitis aguda en el sur de España - debe ser objeto de estudios posteriores. Curiosamente, la distribución geográfica de los casos de hepatitis aguda por fiebre Q en España coincide con el área de distribución de garrapatas del género Hyalomma y estas garrapatas utilizan principalmente a los animales silvestres como hospedadores y ocasionalmente pican a humanos.

Los resultados obtenidos en este estudio podrían ayudar a comprender mejor la epidemiología de C. burnetii en España y el papel de la fauna silvestre y sus garrapatas en la transmisión del patógeno en la interfaz silvestre-humano. Estos resultados dan pie al diseño de estudios enfocados a comprender qué hecho está detrás de la separación geográfica evidente en la presentación clínica aguda de los casos de fiebre Q en humanos en España. Esa información mejoraría los protocolos de prevención de fiebre Q en humanos en nuestro país, lo que podría suponer una reducción en los costes de hospitalización y pérdidas laborales ocasionados por la enfermedad.

La infección por Coxiella burnetii podría suponer pérdidas reproductivas en las

poblaciones de ciervo

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Estimar los efectos clínicos de las infecciones por patógenos que cursan con fallo reproductivo es complejo en la fauna silvestre por la dificultad de acceder a muestras ocasionadas tras el fallo reproductivo (anejos fetales, fetos). Quizás por esta razón el efecto de estas enfermedades, incluida la fiebre Q, haya pasado desapercibido en las poblaciones de animales silvestres a pesar de que existen evidencias de fallo reproductivo asociado a la infección por C. burnetii en numerosas especies de mamíferos silvestes en colecciones zoológicas. Estimar el peso de la fiebre Q en las pérdidas productivas en animales silvestres producidos con interés cinegético, bien en fincas privadas bien en granjas, permitiría evaluar la necesidad de controlar la infección por este patógeno.

En esta Tesis Doctoral se evaluó la implicación de la infección por C. burnetii en un episodio de fallo reproductivo en una población de ciervo producida en condiciones controladas (CAPÍTULO III.1). Desafortunadamente el acceso a fetos, neonatos y anejos fetales no fue posible por las particularidades de producción del ciervo en la explotación que vetan el acceso a las parcelas donde las hembras paren a sus crías. Por ello, para aproximar si algún patógeno asociado a fallo reproductivo podría ser la causa del problema se tomaron muestras de sangre y hisopos vaginales de ciervas que experimentaron fallo reproductivo y de ciervas que parieron. Se analizó la presencia y prevalencia de algunos patógenos reproductivos que se han diagnosticado en casos de fallo reproductivo en rumiantes, incluyendo las bacterias Chlamydia abortus y C. burnetii, y los protozoos Toxoplasma gondii y Neospora caninum. Algunos otros patógenos importantes en el fallo reproductivo, como Brucella spp., no se analizaron al haberse demostrado previamente que esta población era libre. Los resultados de los análisis serológicos llevados a cabo y de los estudios de diagnóstico molecular en secreciones vaginales señalaron la implicación de C. burnetii en el episodio de fallo reproductivo en la población. Sin embargo, la causalidad de la infección por C. burnetii

271 detectada no pudo ser confirmada debido al difícil acceso a las muestras necesarias para el diagnóstico de fallo reproductivo.

Estos resultados indican que se debe advertir a los productores de ciervo de que extremen la vigilancia de los animales durante la época de cría para realizar una toma de muestras adecuada y diagnosticar las causas del fallo reproductivo. Conocer las causas permitirá establecer estrategias que estén enfocadas a reducir dichas pérdidas y las consecuencias económicas de las mismas.

Las vías de excreción de Coxiella burnetii en fauna silvestre son las mismas que las

descritas en rumiantes domésticos, incluyendo

secreciones vaginales, semen, leche y heces

Estimar cuáles son las vías por las que C. burnetii es eliminada por animales infectados es la única vía de conocer cómo se produce la contaminación ambiental por esta bacteria y, por ende, su transmisión a individuos susceptibles. Además, conocer estas vías es la

única forma posible de evaluar el efecto de cualquier estrategia de control de C. burnetii en fauna silvestre y estimar su eficacia.

En los diversos estudios realizados en esta Tesis Doctoral con diferentes especies de fauna silvestre (CAPÍTULOS II.2, III.1, III.2 y IV) en las que se han tomado muestras diversas para estudio de excreción se ha encontrado ADN de C. burnetii en hisopos vaginales/uterinos de ciervo rojo y conejo de monte, en semen de jabalí, en leche/glándula mamaria de ciervo y en heces/hisopos rectales de ciervo y jabalí. En jabalí también se detectó la presencia de ADN de C. burnetii en hisopos nasales, probablemente indicativo de inhalación de aerosoles contaminados con la bacteria. La detección de C. burnetii en las mismas secreciones/excreciones que han sido descritas en rumiantes domésticos sugiere que la vía vaginal, la leche y las heces son rutas de excreción de C. burnetii en

272 especies silvestres infectadas. La presencia de C. burnetii en semen de jabalí sugiere potencial transmisión vaginal en fauna silvestre, aunque esto también puede ocurrir en rumiantes domésticos y silvestres.

Estos resultados señalan que probablemente la contaminación ambiental en ambientes silvestres con C. burnetii se produzca a partir de la excreción de la bacteria en secreciones vaginales y heces. Las bacterias excretadas de esta forma serían las principales responsables de la contaminación ambiental y la transmisión por aerosoles - supuesta aunque aún no probada en fauna silvestre - a otros individuos.

La implementación de vacunación con vacunas inactivadas de fase I podría

controlar la contaminación ambiental y la presión de infección por C. burnetii en poblaciones endémicas de ciervo rojo tras una aplicación prolongada en el tiempo

Los resultados de los estudios epidemiológicos llevados a cabo en esta Tesis Doctoral

(CAPÍTULOS II.1 y II.2) muestran una amplia distribución geográfica de C. burnetii en la fauna silvestre en España con niveles de seroprevalencia similares a los descritos en rumiantes domésticos en el país y que constituyen la principal fuente de infección para el ser humano. Además, algunas de las pocas cepas de C. burnetii de fauna silvestre tipadas también han sido descritas en animales domésticos y personas (CAPÍTULOS II.4 y II.5).

Esta situación muestra la necesidad de diseñar y evaluar estrategias que pueden servir para controlar de forma eficiente la contaminación ambiental por C. burnetii y, con ello, su transmisión. El ciervo rojo podría ser uno de los reservorios silvestres más importantes para C. burnetii en Europa según nuestros resultados. Evaluar la eficacia en el ciervo rojo de vacunas que han demostrado potencial para el control de C. burnetii en especies de rumiantes domésticos proporcionaría una información muy valiosa y, en caso de ser efectiva, una herramienta muy interesante para controlar la infección en poblaciones de

273 ciervo en condiciones controladas. Los resultados de la eficacia de la vacunación podrían ser también extrapolados para adaptar el protocolo vacunal a poblaciones de ciervos en libertad en el futuro.

Así, el último estudio incluido en esta Tesis Doctoral (CAPÍTULO IV) evalúa la eficacia de un protocolo de vacunación de ciervo rojo con una vacuna comercial inactivada de C. burnetii en fase I que ha demostrado eficacia en rumiantes domésticos, que ha sido aprobada por la Agencia Europea del Medicamento para su uso en mamíferos y que está disponible comercialmente. Se diseñó un protocolo de vacunación en fases de manera que en una fase inicial se vacunase a las hembras adultas de la población independientemente de su estado en relación a la infección por C. burnetii y progresivamente se fuese inmunizando a los individuos jóvenes de la explotación antes de su primera exposición a la infección natural. El diseño se hizo con la intención de simular los protocolos que deberían ser diseñados para la implementación de la vacunación en poblaciones de ciervo en libertad contra C. burnetii. Se vacunaron así tres cohortes de ciervas, una cohorte de hembras vacunadas por primera vez a la edad de 2 años o más y 2 cohortes de ciervas vacunadas desde los 13 meses de edad. En cada cohorte se vacunó aproximadamente al

75% de los animales mientras el resto se dejaron sin vacunar constituyendo el grupo control. Para evaluar la capacidad inmunomoduladora de la vacunación y su eficacia sobre la excreción de C. burnetii se tomaron muestras de sangre, hisopos vaginales, leche y heces a diferentes tiempos de la vacunación y durante 3-4 años tras el inicio del experimento. Se estimó la seroconversión por la vacunación en hembras seronegativas antes de la vacunación y la evolución de la inmunidad humoral con el tiempo tras la vacunación. Mediante qPCR se analizó el efecto de la vacunación en el tiempo sobre la excreción de C. burnetii en secreciones vaginales, leche y heces en los grupos vacunal y control. A pesar de la eficacia de la vacuna en la inducción de respuesta inmunológica

274 humoral en las ciervas, ni la prevalencia de excreción ni la cantidad de bacterias excretadas disminuyó en secreciones vaginales y leche con el tiempo de manera significativa en el grupo vacunal con respecto al grupo control. Sin embargo, en ambos grupos se observó una disminución progresiva de la prevalencia de excretores en heces con el tiempo, aunque no de la cantidad excretada. Este último resultado junto con observaciones previas en esta Tesis Doctoral (CAPÍTULO II.3) sobre la disminución de la presión de infección en el rebaño en ciervas jóvenes desde el inicio de la vacunación sugiere que quizás la vacuna pueda tener un efecto a largo plazo sobre la contaminación ambiental. De confirmarse este resultado, las heces se confirmarían como la fuente principal de contaminación ambiental con C. burnetii en ciervo rojo.

Este es el primer trabajo científico que evalúa la eficacia de la aplicación de vacunas comerciales inactivadas de C. burnetii en fase I en ciervo rojo. Al igual que se ha observado en experimentos similares en rumiantes domésticos, parece recomendable aplicar los programas vacunales durante un tiempo más o menos prolongado, al menos superior a 3 años, para observar algún efecto sobre el rebaño. Las dificultades asociadas al coste de las vacunas y al seguimiento de la vacunación no han permitido realizar un seguimiento más prolongado en el tiempo, por lo que este hecho debe ser tenido en cuenta para el diseño de experimentos de vacunación en condiciones de campo en el futuro. Aún así, los resultados son prometedores y señalan que estas vacunas podrían ser un buen método de control de C. burnetii, quizás con mayor eficacia en poblaciones en libertad en las que las prevalencias de infección por C. burnetii son más bajas que en la población objeto de este estudio. Los protocolos futuros de vacunación en ciervo deberían iniciar la vacunación de los animales a edades más tempranas cuando los anticuerpos maternales comienzan a desaparecer y antes de que los animales se enfrenten a la época principal de excreción de C. burnetii por parte de las ciervas gestantes (CAPÍTULO II.3). El diseño

275 del protocolo de vacunación para este estudio se realizó previamente al análisis de la dinámica de infección por C. burnetii en esta misma población (CAPÍTULO II.3), razón por la cual no se pudieron aplicar todas las evidencias de dicho estudio al programa vacunal. A pesar de que la inducción de inmunidad humoral parece durar algo más que la inmunidad humoral inducida por infección natural (CAPÍTULO II.3), quizás sería recomendable plantear protocolos con revacunación cada 6 meses de las hembras que sean seleccionadas para reproducción. En poblaciones naturales será necesario en el futuro evaluar el efecto de la vacuna sobre los machos, ya que estos podrían estar también excretando C. burnetii en heces y contaminar el ambiente.

En conclusión, los trabajos realizados en esta Tesis Doctoral constituyen un documento

único sobre aspectos básicos de la epidemiología y el control de C. burnetii en la fauna silvestre que permitirán profundizar en el conocimiento de la ecología de este patógeno zoonótico en el futuro y mejorar las capacidades de las sociedades humanas para prevenir y controlar el riesgo asociado a esta bacteria. Debemos profundizar más allá de la punta del iceberg que hoy en día constituye el conocimiento existente sobre C. burnetii en la fauna silvestre.

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Conclusiones

1.- Coxiella burnetii está ampliamente distribuida en las poblaciones de ciervo rojo

(Cervus elaphus) y conejo de monte (Oryctolagus cuniculus) en la península ibérica, las cuales presentan niveles de seroprevalencia similares a los descritos en explotaciones de rumiantes domésticos - vacas, cabras y ovejas - en la Península.

Coxiella burnetii is widely distributed in red deer (Cervus elaphus) and European wild rabbit (Oryctolagus cuniculus) populations in the Iberian Penisula, in which it presents seroprevalence levels alike those reported in domestic ruminant - cattle, goat and sheep - herds in the same region.

2.- El ciervo rojo (Cervus elaphus) y el conejo de monte (Oryctolagus cuniculus) son reservorios verdaderos de Coxiella burnetii, al menos en la península ibérica, ya que son susceptibles a la infección por Coxiella burnetii, permiten el desarrollo de infecciones sistémicas por esta bacteria y son capaces de excretarla a través de diferentes secreciones y excreciones permitiendo la transmisión de Coxiella burnetii a otros individuos.

Red deer (Cervus elaphus) and European wild rabbit (Oryctolagus cuniculus) are true reservoirs of Coxiella burnetii, at least in the Iberian Penisula, because they are susceptible to infection by Coxiella burnetii, suffer from systemic infections and shed the bacterium in several secretions and excretions that allow transmission of Coxiella burnetii to other hosts.

3.- En poblaciones de ciervo rojo (Cervus elaphus) en las que Coxiella burnetii es endémica, la seroprevalencia fluctúa en el tiempo en ciervas adultas y jóvenes. Estos cambios podrían estar asociados con la variación observada en la presión de infección y ofrecerían ventanas temporales para introducir programas vacunales que alcancen a una mayor proporción de animales susceptibles en la población.

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In red deer (Cervus elaphus) populations with endemic circulation of Coxiella burnetii, seroprevalence fluctuates in time both in adult and yearling hinds. That changes could be linked to the observed variation in infection pressure and could constitute time windows in which introducing vaccination protocols that would reach a higher proportion of susceptible individuals in the population.

4.- El efecto de la edad en las poblaciones de ciervo rojo (Cervus elaphus) endémicas sobre la respuesta inmunológica humoral frente a la infección por Coxiella burnetii está probablemente vinculado a una mayor capacidad inmunológica de los individuos con la edad.

Age-related effects on the immune humoral response against infection by Coxiella burnetii in endemic red deer (Cervus elaphus) populations are most probably linked to increasing immune capacity of individuals with age.

5.- La vida media de los anticuerpos producidos en ciervo rojo (Cervus elaphus) frente a la infección natural por Coxiella burnetii es de alrededor de 6 meses, aunque ésta aumenta hasta alrededor de 1 año tras la vacunación con vacunas inactivadas de fase I.

Average half-life of antibodies in red deer (Cervus elaphus) after natural infection by

Coxiella burnetii is around 6 months although it increases up to around 1 year after vaccination with phase I inactivated vaccines.

6.- La excreción de Coxiella burnetii en hembras de ciervo rojo (Cervus elaphus) presenta un patrón estacional claro con predominancia de excreción alrededor de la época de partos. Esto unido a la corta duración de los anticuerpos tras la infección natural conlleva menor seroprevalencia en invierno, lo que implica que los estudios epidemiológicos basados en muestras de animales silvestres recolectadas en época de caza - octubre a febrero - subestimen la seroprevalencia real.

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Coxiella burnetii shedding by red deer (Cervus elaphus) females displays a seasonal pattern with major shedding around the breeding season. That fact in addition to the low average half-life of antibodies after natural infeccion triggers lower seroprevalence in winter and therefore epidemiological studies based on wildlife samples collected during the hunting season - October to February - subestimate real seroprevalence.

7.- Las crías de ciervo rojo (Cervus elaphus) reciben de sus madres una alta carga de anticuerpos en la lactación que desaparecen antes de los 7 meses de vida. Estas cargas elevadas de anticuerpos podrían proporcionar cierta protección frente a la infección por

Coxiella burnetii a estos animales que en su nacimiento están expuestos a altas cargas bacterianas excretadas por las hembras paridas.

Red deer (Cervus elaphus) calves are given high doses of antibodies by their mothers during lactation that disappear before their 7th month of life. That high antibody dose could protect calves from infection by Coxiella burnetii which at birth are exposed to high burdens of infectious bacteria shed by farrowed females.

8.- La fauna silvestre comparte genotipos de Coxiella burnetii con el ganado doméstico, el ser humano y garrapatas.

Widlife shares Coxiella burnetii genotypes with livestock, human beings and ticks.

9.- Los genotipos de Coxiella burnetii que circulan en poblaciones simpátrica de ciervo rojo (Cervus elaphus) y conejo de monte (Oryctolagus cuniculus) se agrupan en función de su hospedador, lo que sugiere que existe algún tipo de adaptación del patógeno a sus hospedadores.

Coxiella burnetii genotypes circulating in sympatric red deer (Cervus elaphus) and

European wild rabbit (Oryctolagus cuniculus) populations cluster according to their host of origin, which suggest that there is some adaptation of the pathogen to its hosts.

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10.- El tipado genético mediante MLVA de cepas de Coxiella burnetii de ciervo rojo

(Cervus elaphus) y conejo de monte (Oryctolagus cuniculus) muestra mayor similitud de los genotipos presentes en la fauna silvestre con genotipos aislados de casos clínicos de fiebre Q en humanos que con los genotipos aislados en ganado. El genotipado mediante

PCR-RLB confirma estos resultados.

MLVA genotyping of Coxiella burnetii strains from red deer (Cervus elaphus) and

European wild rabbit (Oryctolagus cuniculus) shows higher similarities of wildlife genotypes to genotypes isolated from human Q fever clinical cases than to genotypes isolated in livestock. PCR-RLB genotyping confirms that findings.

11.- El genotipado de cepas de Coxiella burnetii en fauna silvestre en España mediante

PCR-RLB señala grandes similitudes con genotipos de garrapatas y con genotipos aislados en casos clínicos agudos de hepatitis en humanos. Estos resultados sugieren que algunos genotipos de Coxiella burnetii son mantenidos en un ciclo que incluye especies silvestres y garrapatas y que estos genotipos podrían ser ocasionalmente transmitidos a seres humanos a través de la picadura de garrapatas o por exposición a fauna silvestre.

PCR-RLB genotyping of Coxiella burnetii strains of wildlife origin in Spain shows high similarities with genotypes of ticks and those isolated from acute clinical cases of hepatitis in humans. That results suggest that certain Coxiella burnetii genotypes are maintained in a cycle that includes wildlife and ticks and that, ocassionally, could be transmitted to humans through tick bites or through exposure to wildlife.

12.- Coxiella burnetii puede ser excretada en ciervo rojo (Cervus elaphus), jabalí (Sus scrofa) y conejo de monte (Oryctolagus cuniculus) en secreciones vaginales, semen, leche y heces.

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Coxiella burnetii can be shed by red deer (Cervus elaphus), Eurasian wild boar (Sus scrofa) and European wild rabbit (Oryctolagus cuniculus) in vaginal secretions, semen, milk and faeces.

13.- La implementación de un programa vacunal frente a Coxiella burnetii basado en vacunas inactivadas de fase I en una población endémica de ciervo rojo (Cervus elaphus) no reduce ni la prevalencia de excreción ni la cantidad de bacterias excretadas en secreciones vaginales ni en leche en comparación con el grupo control, aunque sí se observa una reducción en la prevalencia de excretores de Coxiella burnetii en heces en el tiempo tanto en el grupo vacunal como en el grupo control en simpatría. Este hecho unido a una disminución en la incidencia de infección por Coxiella burnetii en hembras jóvenes en la población tras la implementación de la vacunación sugiere que quizás a largo plazo la vacuna tenga un efecto sobre la reducción de la contaminación ambiental por Coxiella burnetii en poblaciones de ciervo rojo.

The implementation of a vaccination program against Coxiella burnetii based upon inactivated phase I vaccines in an endemic red deer (Cervus elaphus) population does not account in a reduction of shedding prevalence and bacterial burden in vaginal secretions and milk in comparison to the control group; however a reduction in the prevalence of

Coxiella burnetii shedders in faeces with time both in the vaccinated and the sympatric control group is observed. This fact together with the observed decreasing incidence of

Coxiella burnetii infection in yearling hinds in the population after the implementation of vaccination suggest that perhaps in the long-time scale vaccination would reduce environmental contamination with Coxiella burnetii in red deer populations.

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Bibliografía Acevedo P, Delibes-Mateos M, Escudero MA, Vicente J, Marco J, Gortazar C. 2005.

Environmental constraints in the colonization sequence of roe deer (Capreolus capreolus

Linnaeus, 1758) across the Iberian Mountains, Spain. Journal of Biogeography. 32(9):

1671-1680

Acevedo P, Ferreres J, Jaroso R, Durán M, Escudero MA, Marco J, Gortázar C. 2010a.

Estimating roe deer abundance from pellet group counts in Spain: An assessment of methods suitable for Mediterranean woodlands. Ecological Indicators. 10(6): 1226-1230

Acevedo P, Ruiz-Fons F, Estrada R, Márquez AL, Miranda MA, Gortázar C, Lucientes

J. 2010b. A broad assessment of factors determining Culicoides imicola abundance: modelling the present and forecasting its future in climate change scenarios. PLoS One

5:e14236.

Acevedo P, Ruiz-Fons F, Vicente J, Reyes-García AR, Alzaga V, Gortázar C. 2008.

Estimating red deer abundance in a wide range of management situations in

Mediterranean habitats. J Zool. 276:37–47.

Acevedo P, Vicente J, Höfle U, Cassinello J, Ruiz-Fons F, Gortazar C. 2007. Estimation of European wild boar relative abundance and aggregation: A novel method in epidemiological risk assessment. Epidemiol Infect. 135(3): 519-527

Acevedo P, Quirós-Fernández F, Casal J, Vicente J. 2014. Spatial distribution of wild boar population abundance: basic information for spatial epidemiology and wildlife management. Ecol Indic 36:594– 600. doi:10.1016/j.ecolind.2013.09.019

Agerholm JS. 2013. Coxiella burnetii associated reproductive disorders in domestic animals-a critical review. Acta Vet Scand. 55:13

282

Agerholm JS. 2014. Coxiella burnetii and reproductive disorders in cattle. Infect Genet

Evol. 28: 150-150

Akaike H. 1974. A new look at the statistical model identification. Institute of Electrical and Electronics Engineers Transactions on Automatic Control. 19:716–725 de Alarcón A, Villanueva JL, Viciana P, López-Cortés L, Torronteras R, Bernabeu M,

Cordero E, Pachón J. 2003. Q fever: Epidemiology, clinical features and prognosis. A study from 1983 to 1999 in the South of Spain. J Infection. 47(2): 110-116

Alexander KA, McNutt JW. 2010. Human behavior influences infectious disease emergence at the human-animal interface. Front Ecol Environ. 8(10): 522-526

Almeida AP, Marcilia A, Leite RC, Nieri-Bastosa FA, Domingues LN, Martins JR,

Labruna MB. 2012. Coxiella symbiont in the tick Ornithodoros rostratus (Acari:

Argasidae). Ticks Tick Borne Dis. 3: 203-206

Alonso E, Lopez-Etxaniz I, Hurtado A, Liendo P, Urbaneja F, Aspiritxaga I, Olaizola JI,

Piñero A, Arrazola I, Barandika JF, Hernáez S, Muniozguren N, García-Pérez AL. 2015.

Q Fever Outbreak among Workers at a Waste-Sorting Plant. PLoS One. 10(9):e0138817.

Álvarez J, Pérez A, Mardones FO, Pérez-Sancho M, García-Seco T, Pagés E, Mirat F,

Díaz R, Carpintero J, Domínguez L. 2012. Epidemiological factors associated with the exposure of cattle to Coxiella burnetii in the Madrid region of Spain. Vet J. 194:102–107.

Amitai Z, Bromberg M, Bernstein M, Raveh D, Keysary A, David D, Pitlik S, Swerdlow

D, Massung R, Rzotkiewicz S, Halutz O, Shohat T. 2010. A large Q fever outbreak in an urban school in Central Israel. Clin Infect Dis. 50(11): 1433-1438

283

Andoh, M., Nagaoka, H., Yamaguchi, T., Fukushi, H., Hirai, K., 2004. Comparison of

Japanese isolates of C. burnetii by PCR-RFLP and sequence analysis. Microbiol.

Immunol. 48:971-975

Angelakis E & Raoult D. 2010. Q fever. Vet Microbiol 140:297-309.

Anheyer-Behmenburg HE. 2013. Untersuchungen zum Vorkommen von

Zoonoseerregern und dem kaninen Staupevirus in der Waschbärpopulation

Niedersachsens, 2011-2013. PhD Thesis

Anne-Frieda Taurel, Guatteo R, Joly A, Beaudeau F. 2012. Effectiveness of vaccination and antibiotics to control Coxiella burnetii shedding around calving in dairy cows. Vet

Microbiol. 159: 432–437

Apanaskevich DA, Santos-Silva MM, Horak IG. The genus Hyalomma Koch, 1844. IV.

Redescription of all parasitic stages of H. (Euhyalomma) lusitanicum Koch, 1844 and the adults of H. (E.) franchinii Tonelli Rondelli, 1932 (Acari: Ixodidae) with a first description of its immature stages. 2008. Folia Parasitologica 55: 61–74

Apollonio M, Andersen R, Putman R. 2010. European ungulates and their management in the 21st century. Cambridge University Press, Cambridge, United Kingdom.

Arricau-Bouvery N, Souriau A, Lechopier P, Rodolakis A. 2003. Excretion of Coxiella burnetii during an experimental infection of pregnant goats with an abortive goat strain

CbC1. Ann N Y Acad Sci. 990: 524-526

Arricau-Bouvery N & Rodolakis A. 2005. Is Q fever an emerging or re-emerging zoonosis? Vet Res. 36(3): 327-349

284

Arricau-Bouvery N, Souriau A, Bodier C, Dufour P, Rousset E, Rodolakis A. 2005. Effect of vaccination with phase I and phase II Coxiella burnetii vaccines in pregnant goats.

Vaccine. 23: 4392-4402.

Arricau-Bouvery N, Hauck Y, Bejauoi A, Frangoulidis D, Bodier CC, Souriau A, Meyer

H, Neubauer H, Rodolakis A, Vergnaud G. 2006. Molecular characterization of Coxiella burnetii isolates by infrequent restriction site-PCR and MLVA typing. BMC Microbiol.

6:38

Artois M, Blancou J, Dupeyroux O, Gilot-Fromont E. 2011. Sustainable control of zoonotic pathogens in wildlife: how to be fair to wild animals? Rev Sci Tech Off. Int

Epizoot. 30(3): 733-743

Astobiza I, Aduriz G, Atxaerandio R, Barandika JF, Hurtado A, Povedano I, Juste RA,

García-Pérez AL. 2007. Estimacion de la prevalencia de Coxiella burnetii en ganado ovino por metodos serologicos y moleculares. XII Simposio de AVEDILA. Bilbao.

España

Astobiza I, Barandika JF, Hurtado A, Juste RA, Garcia- Perez AL. 2010. Kinetics of

Coxiella burnetii excretion in a commercial dairy sheep flock after treatment with oxytetracycline. Vet. J. 184:172–175

Astobiza I, Barral M, Ruiz-Fons F, Barandika JF, Gerrikagoitia X, Hurtado A, García-

Pérez AL. 2011a. Molecular investigation of the occurrence of Coxiella burnetii in wildlife and ticks in an endemic area. Vet Microbiol. 147: 190-194

Astobiza I, Barandika JF, Ruiz-Fons F, Hurtado A, Povedano I, Juste RA, García-Pérez

AL. 2011b. Coxiella burnetii shedding and environmental contamination at lambing in two highly naturally-infected dairy sheep flocks after vaccination. Res Vet Sci. 91: e58- e63. DOI:10.1016/j.rvsc.2010.11.014

285

Astobiza I, Barandika JF, Ruiz-Fons F, Hurtado A, Povedano I, Juste RA, García-Pérez

AL. 2011c. Four-year evaluation of the effect of vaccination against Coxiella burnetii on reduction of animal infection and environmental contamination in a naturally infected dairy sheep flock. Appl Environ Microbiol. 77:7405-7407

Astobiza I, Ruiz-Fons F, Piñero A, Barandika JF, Hurtado A, García- Pérez AL. 2012a.

Estimation of Coxiella burnetii prevalence in dairy cattle in intensive systems by serological and molecular analyses of bulk-tank milk samples. J Dairy Sci. 95: 1632-

1638.

Astobiza I, Tilburg JJHC, Piñero A, Hurtado A, García-Pérez AL, Nabuurs-Franssen MH,

Klaassen CHW. 2012b. Genotyping of Coxiella burnetii from domestic ruminants in northern Spain. BMC Vet Res. 8: 241

Astobiza I, Barandika JF, Juste RA, Hurtado A, García-Pérez AL. 2013. Evaluation of the efficacy of oxytetracycline treatment followed by vaccination against Q fever in a highly infected sheep flock. Vet J. 196(1): 81-85

Aulagnier S, Giannatos G, Herrero J. 2008a. Rupicapra rupicapra. The IUCN Red List of Threatened Species 2008: e.T39255A10179647. http://dx.doi.org/10.2305/IUCN.UK.2008.RLTS.T39255A10179647.en . Downloaded on 04 November 2015.

Aulagnier S, Kranz A, Lovari S, Jdeidi T, Masseti M, Nader I, de Smet K, Cuzin F. 2008b.

Capra ibex. The IUCN Red List of Threatened Species 2008: e.T42397A10695445. http://dx.doi.org/10.2305/IUCN.UK.2008.RLTS.T42397A10695445.en . Downloaded on 04 November 2015

Babudieri B. 1959. Q fever: a zoonosis. Adv. Vet. Sci. 5:81.

286

Baca OG & Paretsky D. 1983. Q fever and Coxiella burnetii: a model for host-parasite interactions. Microbiol Rev. 47:127-149.

Balasch M, Pujols J, Segalés J, Plana-Durán J, Pumarola M. 1998. Study of the persistence of Aujeszky’s disease (pseudorabies) virus in peripheral blood mononuclear cells and tissues of experimentally infected pigs. Vet Microbiol. 62:171–183.

Banazis MJ, Bestall AS, Reida SA, SG Fenwick. 2010. A survey of Western Australian sheep, cattle and kangaroos to determine the prevalence of Coxiella burnetii. Vet

Microbiol. 143: 337-345

Baneth G. 2014. Tick-borne infections of animals and humans: A common ground.

International Journal for Parasitology. 44(9): 591-596

Baradel JM, Barrat J, Blancou J, Boutin JM, Chastel C, Dannacher G, Delorme D, Gerard

Y, Gourreau JM, Kihm U, Larenaudie B, Le Goff C, Pastoret PP, Perreau P, Schwers A,

Thiry E, Trap D, Uilenberg G, Vannier P. 1988. Results of a serological survey of wild mammals in FranceRev. sci. tech. Off int Epiz. 7(4): 873-883.

Barandika J, Hurtado A, García-Esteban C, Gil H, Escudero R, Barral M, Jado I, Juste R,

Anda P, García-Perez AL. 2007. Tick-borne zoonotic bacteria in wild and domestic small mammals in Northern Spain. Appl Environ Microbiol. 73:6166–6171

Barandika, J.F, Hurtado, A, García-Sanmartín, J, Juste, R.A, Anda, P, García-Pérez, A.L.

2008. Prevalence of tick-borne zoonotic bacteria in questing adult ticks from Northern

Spain. 2008. Vector-Borne and Zoonotic Diseases. 8 (6):829-835

Beare PA, Samuel JE, Howe D, Virtaneva K, Porcella SF, Heinzen RA. 2006. Genetic diversity of the Q fever agent, Coxiella burnetii, assessed by microarray-based whole- genome comparisons. J Bacteriol 188: 2309-2324

287

Beltrán-Beck B, Ballesteros C, Vicente J, De La Fuente J, Gortázar C. 2012a. Progress in oral vaccination against tuberculosis in its main wildlife reservoir in Iberia, the Eurasian wild boar. Veterinary Medicine International. 2012, 978501

Beltrán-Beck B, García FJ, Gortázar, C. 2012b. Raccoons in Europe: Disease hazards due to the establishment of an invasive species. European Journal of Wildlife Research.

58 (1): 5-15

Beltrán-Beck B, De La Fuente J, Garrido JM, Aranaz A, Sevilla I, Villar M, Boadella M,

Galindo RC, Pérez De La Lastra JM, Moreno-Cid JA, Fernández De Mera IG, Alberdi P,

Santos G, Ballesteros C, Lyashchenko KP, Minguijón E, Romero B, De Juan L,

Domínguez L, Juste R, Gortazar C. 2014. Oral vaccination with heat inactivated

Mycobacterium bovis activates the complement system to protect against tuberculosis.

PLoS ONE. 9(5): e98048

Berri M, Laroucau K, Rodolakis A. 2000. The detection of Coxiella burnetii from ovine genital swabs, milk and fecal samples by the use of a single touchdown polymerase chain reaction. Vet Microbiol. 72:285-293

Berri M, Souriau A, Crosby M, Rodolakis A. 2002. Shedding of Coxiella burnetii in ewes in two pregnancies following an episode of Coxiella abortion in a sheep flock. Veterinary

Microbiology. 85(1): 55-60

Berri M, Souriau A, Crosby M, Crochet D, Lechopier P, Rodolakis A. 2001.

Relationships between the shedding of Coxiella burnetii, clinical signs and serological responses of 34 sheep. Vet Rec. 148: 502-505.

Berri M, Rousset E, Hechard C, Champion JL, Dufour P, Russo P, Rodolakis A. 2005.

Progression of Q fever and Coxiella burnetii shedding in milk after an outbreak of enzootic abortion in a goat herd. Vet Rec. 156(17): 548-549

288

Berri M, Rousset E, Champion JL, Russo P, Rodolakis A. Goats may experience reproductive failures and shed Coxiella burnetii at two successive parturitions after a Q fever infection. Research in Veterinary Science. 83(1): 47-52

Bewley KR. 2013. Animal models of Q fever (Coxiella burnetii). Comparative Med.

63(6): 469-476

Biesiada G, Czepiel J, Leśniak MR, Garlicki A, Mach T. 2012. Lyme disease: Review.

Archives of Medical Science. 8(6): 978-982

Bjork A, Marsden-Haug N, Nett RJ, Kersh GJ, Nicholson W, Gibson D, Szymanski T,

Emery M, Kohrs P, Woodhall D, Anderson AD. 2014. First reported multistate human Q fever outbreak in the United States, 2011. Vector-Borne Zoonotic Dis. 14(2): 111-117

Blancou J. 1983. Serologic testing of wild roe deer (Capreolus capreolus) from the Trois

Fontaines Forest Region of Eastern France. J Wildlife Dis. 19(3):271-273

Boadella M, Acevedo P, Vicente J, Mentaberre G, Balseiro A, Arnal MC, Martínez D,

García-Bocanegra I, Casal C, Álvarez J, Oleaga A, Lavín S, Muñoz M, Sáez- Llorente

JL, de la Fuente J, Gortázar C. 2011. Spatio-temporal trends of Iberian wild boar contact with Mycobacterium tuberculosis complex detected by ELISA. EcoHealth 8:478–484.

Boadella M, Carta T, Oleaga A, Pajares G, Muñoz M, Gortázar C. 2010. Serosurvey for selected pathogens in Iberian roe deer. BMC Vet Res 6:51.

Boadella M, Gortázar C, Vicente J, Ruiz-Fons F. 2012a. Wild boar: an increasing concern for Aujeszky's disease control in pigs? BMC Vet Res. 8:7.

Boadella M, Ruiz-Fons JF, Vicente J, Martín M, Segalés J, Gortázar C. 2012.

Seroprevalence evolution of selected pathogens in Iberian wild boar. Transbound

Emerg. Dis. 59:395–404.

289

Boadella M, Vicente J, Ruiz-Fons F, de la Fuente J, Gortázar C. 2012. Effects of culling

Eurasian wild boar on the prevalence of Mycobacterium bovis and Aujeszky's disease virus. Prev Vet Med. 107:214–221

Böttcher J, Vossen A, Janowetz B, Alex M, Gangl A, Randt A, Meier N. 2011. Insights into the dynamics of endemic Coxiella burnetii infection in cattle by application of phase- specific ELISAs in an infected dairy herd. Vet Microbiol. 151: 291-300

Brezina R. 1958. Contribution to the study of phase variation in Coxiella burnetii. Acta virol. 2:91- 102.

Brom RD, van Engelen E, Roest HIJ, Hoek WD, Vellema P. 2015. Coxiella burnetii infections in sheep or goats: An opinionated review. Vet Microbiol. In press

Brooke RJ, Kretzschmar MEE, Mutters NT, Teunis PF. Human dose response relation for airborne exposure to Coxiella burnetii. 2013. BMC Infect Dis. 13(1):488

Brooke RJ, Mutters NT, Péter O, Kretzschmar MEE, Teunis PFM. 2015. Exposure to low doses of Coxiella burnetii caused high illness attack rates: Insights from combining human challenge and outbreak data. Epidemics. 11:1-6

Burgdorfer W, Pickens EG, Newhouse VF, Lackman DB. 1963. Isolation of Coxiella burnetii from rodents in western Montana. J Infect Dis. 112:181-186

Burnham KP, Anderson DR. 2002. Model selection and multi-model inference. Springer,

New York, NY.

Cabello J, Altet L, Napolitano C, Sastre N, Hidalgo E, Dávila JA, Millán J. 2013. Survey of infectious agents in the endangered Darwin’s fox (Lycalopex fulvipes): High prevalence and diversity of hemotrophic mycoplasmas. Vet Microbiol. 167: 448-454

290

Candela MG, Serrano E, Sevila J, León L, Caro MR, Verheyden H. 2014. Pathogens of zoonotic and biological importance in roe deer (Capreolus capreolus): Seroprevalence in an agro-system population in France. Res Vet Sci. 96: 254-259

Capuano F, Perugini AG, Parisi A, Montagna CO, Nilvelli M. 2004. Improved detection of Coxiella burnetii in cows milk by immunomagnetic separation and PCR. Vet Res

Commun. 28: 279-282.

Caron A, Miguel E, Gomo C, Makaya P, Pfukenyi DM, Foggin C, Hove T, de Garine-

Wichatitsky M. 2013. Relationship between burden of infection in ungulate populations and wildlife/livestock interfaces. Epidemiol Infect. 141(7): 1522-1535.

Carstensen M, O’Brien DJ, Schmitt SM. 2011. Public acceptance as a determinant of management strategies for bovine tuberculosis in free-ranging U.S. wildlife. Vet

Microbiol. 151(1–2):200–204

Chaber AL, Lloyd C, O’Donovan D, McKeown S, Wernery U, Bailey T. 2012. A

Serologic Survey for Coxiella burnetii in Semi-wild Ungulates in the Emirate of Dubai,

United Arab Emirates. Journal of Wildlife Diseases. 48(1): 220-222

Chmielewski T, Sidi-Boumedine K, Duquesne V, Podsiadly E, Thiéry R, Tylewska-

Wierzbanowska S. 2009. Molecular epidemiology of Q fever in Poland. Pol J Microbiol.

58: 9-13

Chomel BB, Carniciu ML, Kasten RW, Castelli PM, Work TM, Jessup DA. 1994.

Antibody prevalence of eight ruminant infectious diseases in California mule and black- tailed deer (Odocoileus Hemionus). J Wildlife Dis. 30(1):51-59

Clark RK, Jessup DA, Hird DW, Ruppanner R, Meyer ME. 1983. Serologic survey of

California wild hogs for antibodies against selected zoonotic disease agents. J Am Vet

Med Assoc. 183(11): 1248-1251

291

Clemente L, Fernandes TL, Barahdna R, Bernardino R, Botelho A. 2008. Confirmation by PCR of Coxiella burnetii infection in animals at a zoo in Lisbon, Portugal. Vet Rec.

163: 221-222

Clutton-Brock TH, Guinness FE, Albon SP. 1982. Red deer: behaviour and ecology of two sexes. University of Chicago Press, Chicago, IL.

Cook MJ. 2014. Lyme borreliosis: A review of data on transmission time after tick attachment. Int J Gen Med. 8: 1-8

Coop RL & Kyriazakis I. 1999. Nutrition-parasite interaction. Vet Parasitol. 84: 187-204

Coop RL & Kyriazakis I. 2001. Influence of host nutrition on the development and consequences of nematode parasitism in ruminants. Trends Parasitol. 17: 325-330

Cooper A, Hedlefs R, McGowan M, Ketheesan N, Govan B. 2011. Serological evidence of Coxiella burnetii infection in beef cattle in Queensland. Aust Vet J. 89: 260-264

Cooper A, Stephens J, Ketheesan N, Govan B. 2013. Detection of Coxiella burnetii DNA in Wildlife and Ticks in Northern Queensland, Australia. Vector Borne Zoonotic Dis. 13

(1): 12-16

Courcoul A, Monod H, Nielen M, Klinkenberg D, Hogerwerf L, Beaudeau F, Vergu E.

2011. Modelling the effect of heterogeneity of shedding on the within herd Coxiella burnetii spread and identification of key parameters by sensitivity analysis. J Theor Biol.

284(1): 130-141

Cowled BD, Giannini F, Beckett SD, Woolnough A, Barry S, Randall L, Garner G. 2009.

Feral pigs: predicting future distributions. Wildl Res. 36:242–251.

292

Cumbassá A, Barahona MJ, Cunha MV, Azórin B, Fonseca C, Rosalino LM, Tilburg J,

Hagen F, Santos AS, Botelho A. 2015. Coxiella burnetii DNA detected in domestic ruminants and wildlife from Portugal. Veterinary Microbiology. 180(1-2): 136-141

D'amato F, Million M, Edouard S, Delerce J, Robert C, Marrie T, Raoult D. 2014. Draft genome sequence of Coxiella burnetii Dog Utad, a strain isolated from a dog-related outbreak of Q fever. New Microbes New Infect. 2: 136-137

Darwich L, Cabezón O, Echeverria I, Pabón M, Marco I, Molina-López R, Alarcia-Alejos

O, López-Gatius F, Lavín S, Almería S. 2012. Presence of Toxoplasma gondii and

Neospora caninum DNA in the brain of wild birds. Vet. Parasitol. 183:377–381.

Daszak P, Cunningham AA, Hyatt AD. 2000. Emerging infectious diseases of wildlife - threats to biodiversity and human health. Science. 287: 443-449

Daszak P, Zambrana-Torrelio C, Bogich TL, Fernandez M, Epstein JH, Murray KA,

Hamilton H. 2013. Interdisciplinary approaches to understanding disease emergence: The past, present, and future drivers of Nipah virus emergence. Proceedings of the National

Academy of Sciences of the United States of America. 110(1): 3681-3688

Davis KF & D'Odorico P. 2015. Livestock intensification and the influence of dietary change: A calorie-based assessment of competition for crop production. Sci Total

Environ. 538: 817-823

Davoust B, Marié JL, Pommier de Santi V, Berenger JM, Edouard S, Raoult D. 2014.

Three-toed sloth as putative reservoir of Coxiella burnetii, Cayenne, French Guiana.

Emerg Infect Dis 2014;20:1760-61.

Dawe KL, Bayne EM, Boutin S. 2014. Influence of climate and human land use on the distribution of white-tailed deer (Odocoileus virginianus) in the western boreal forest.

Can J Zool. 92(4): 353-363

293 de Bruin A, de Groot A, de Heer L, Bok J, Wielinga PR, Hamans M, van Rotterdam B J,

Janse I. 2011. Detection of Coxiella burnetii in complex matrices by using multiplex quantitative PCR during a major Q fever outbreak in The Netherlands. Appl Environ

Microbiol. 77: 6516-6523 de Bruin A, van Alphen PT, van der Plaats RQ, de Heer LN, Reusken CB, van Rotterdam

BJ, Janse I. 2012. Molecular typing of Coxiella burnetii from animal and environmental matrices during Q fever epidemics in the Netherlands. BMC Vet Res. 8:165.

De Cremoux R, Rousset E, Touratier A, Audussea G, Nicollet P, Ribaud D, David V, Le

Pape M. 2012. Coxiella burnetii vaginal shedding and antibody responses in dairy goat herds in a context of clinical Q fever outbreaks. FEMS Immunol Med Microbiol. 64: 120-

122.

Deforge JR, Cone LA. 2006. The serologic prevalence of Q fever (Coxiella Burnetii).

Complement-fixing antibodies in the peninsular bighorn sheep of southern California.

Am J Trop Med Hyg. 75(2): 315-317

Dekker H. 1975. A simple mathematical model of rodent population cycles. J Math Biol.

19. VI. 2:57-67

Delgado CL, Rosegrant MW, Steinfeld H, Ehui SK, Courbois C. 1999. Livestock to 2020:

The next food revolution. IFPRI, Washington DC.

Delgado CL. 2005. Rising demand for meat and milk in developing countries: implications for grasslands-based livestock production. M.G. DA (Ed.), Grassland: A global resource, Wageningen Academic Publishers, Wageningen (The Netherlands). pp.

29-39

Delibes M, Aymerich M, Cuesta L. 1984. Feeding habits of the Egyptian mongoose or ichneumon in Spain. Acta Theriol (Warsz). 29:205-218

294

Delibes-Mateos M, Ferreras P, Villafuerte R. 2008. Rabbit populations and game management: The situation after 15 years of rabbit haemorrhagic disease in central southern Spain. 2008. Biodiversity and Conservation. 17(3): 559-574

Delibes-Mateos M, Farfán MÁ, Olivero J, Márquez AL, Vargas JM. 2009. Longterm changes in game species over a long period of transformation in the Iberian Mediterranean landscape. Environmental Management. 43(6): 1256-1268.

Denison AM, Thompson HA, Massung RF. 2007. IS1111 insertion sequences of C. burnetii: characterization and use for repetitive element PCR-based differentiation of C. burnetii isolates. BMC Microbiol. 7:91.

Derrick EH. 1937. Q fever, new fever entity: clinical features, diagnosis and laboratory investigation. Med J Aust. 2: 281-299

Derrick EH, Smith DJW, Brown HE. 1939. The role of the Bandicoot in the epidemiology of Q fever: a preliminary study. Med J Aust. 1: 150-155

Derrick EH. 1973. The course of infection with Coxiella burnetii. Med J Aust. 1: 1051-

1057.

Díaz-Fernández S, Arroyo B, Casas F, Martinez-Haro M, Viñuela J. 2013. Effect of Game

Management on Wild Red-Legged Partridge Abundance. PLoS ONE. 8(6):e66671

Dobson A & Foufopoulos J. 2001. Emerging infectious pathogens of wildlife. Philos

Trans R Soc Lond B Biol Sci. 356(1411):1001-1012

Dorko E, Rimárová K, Pilipčinec E, Trávniček M. 2009. Prevalence of Coxiella Burnetii antibodies in wild ruminants in Kavečany Zoo, Košice, Eastern Slovakia. Ann Agric

Environ Med. 16:321–324

295

Dorko E, Rimárová K, Pilipčinec E. 2012. Influence of the environment and occupational exposure on the occurrence of Q fever. Central European Journal of Public Health. 20(3):

208-214

Dubey JP, Desmonts G. 1987. Serological responses of equids fed Toxoplasma gondii oocysts. Equine Vet J. 19:337– 339.

Dunbar MR, Cunningham MW, Roof JC. 1998. Seroprevalence of selected disease agents from free-ranging black bears in Florida. J Wildl Dis. 34(3): 612-619

Duncan C, Gill VA, Worman K, Burek-Huntington K, Pabilonia KL, Johnson S,

Fitzpatrick KA, Weller C, Kersh GJ. 2015. Coxiella burnetii exposure in northern sea otters Enhydra lutris kenyoni. Dis Aquat Org. Vol. 114:83–87

Duncan C, Kersh GJ, Spraker T, Patyk KA, Fitzpatrick KA, Massung RF, Gelatt T. 2012.

Coxiella burnetii in northern fur seal (Callorhinus ursinus) placentas from St. Paul Island,

Alaska. Vector Borne Zoonotic Dis. 12(3):192-195

Duncan C, Savage K, Williams M, Dickerson B, Kondas AV, Fitzpatrick KA, Guerrero

JL, Spraker T, Kersh GJ. 2013. Multiple strains of Coxiella burnetii are present in the environment of St. Paul Island, Alaska. Transbound Emerg Dis. 60: 345-350

Duron O, Jourdain E, McCoy KD. 2014. Diversity and global distribution of the Coxiella intracellular bacterium in seabird ticks. Ticks Tick Borne Dis. 5: 557-563

Dupont H, Raoult D, Brouqui P, Janbon F, Peyramond D, Weiller PJ, Chicheportiche C,

Nezri M, Poirier R. 1992. Epidemiologic features and clinical presentation of acute Q fever in hospitalized patients: 323 French cases. Am J Med. 93: 427-434

EFSA. 2010. Panel on Animal Health and Welfare (AHAW); Scientific Opinion on Q

Fever. EFSA J. 8.1595:114.

296

EFSA. 2014a. The European Union summary report on trends and sources of zoonoses, zoonotic agents and food-borne outbreaks in 2012. EFSA J. 12:3547.

EFSA. 2014b. Evaluation of possible mitigation measures to prevent introduction and spread of African swine fever virus through wild boar. EFSA J. 12:3616.

Ejercito CL, Cai L, Htwe KK, Taki M, Inoshima Y, Kondo T, Kano C, Abe S, Shirota K,

Sugimoto T, Yamaguchi T, Fukushi H, Minamoto N, Kinjo T, Isogai E, Hirai K. 1993.

Serological evidence of Coxiella burnetii infection in wild animals in Japan. J Wildl Dis.

29:481–484.

Eldin C, Mahamat A, Djossou F, Raoult D. 2015. Rainfall and Sloth Births in May, Q

Fever in July, Cayenne, French Guiana. Am J Trop Med Hyg. 92(5):979-981

Enright JB, Franti CE, Longhurst WM, Behymer DE, Wright ME, Dutson VJ. 1971.

Coxiella burnetti in a wildlife-livestock environment; antibody response of ewes and lambs in an endemic Q fever area. Amer J Epidem 94: 62-71

Espejo E, Gil-Díaz A, Oteo JA, Castillo-Rueda R, García-Alvarez L, Santana-Báez S,

Bella F. 2014. Clinical presentation of acute Q fever in Spain: Seasonal and geographical differences. Int J Infect Dis. 26: e162-e164

Estrada-Peñ, A. Bouattour, J.-L. Camicas, A.R. Walker. 2004. Ticks of Domestic

Animals in the Mediterranean Region. A Guide to Identification of Species.

Estrada-Peña A, Farkas R, Jaenson TGT, Koenen F, Madder M, Pascucci I, Salman M,

Tarrés-Call J, Jongejan F. Association of environmental traits with the geographic ranges of ticks (Acari: Ixodidae) of medical and veterinary importance in the western Palearctic.

A digital data set. Exp Appl Acarol (2013) 59:351–366

FAO. Food and Agriculture Organization. Trade.

[http://faostat.fao.org/site/604/DesktopDefault.aspx?PageID=604#ancor]

297

Fernández-de-Mera IG, Vicente J, Höfle U, Fons FR, Ortiz JA, Gortázar C. 2009. Factors affecting red deer skin test responsiveness to bovine and avian tuberculin and to phytohaemagglutinin. Preventive Veterinary Medicine. 90(1-2): 119-126

Fernández Guerrero ML. 2014. Q fever in Spain: “An inconclusive history”. Enferm

Infecc Microbiol Clin. 32(4): 211–212

Fernandez-Llario P, Carranza J. 2000. Reproductive performance of the wild boar in a

Mediterranean ecosystem under drought conditions. Ethol Ecol Evol. 12(4): 335-343

Flueck WT, Smith-Flueck JM, Naumann CM. 2003. The current distribution of red deer

(Cervus elaphus) in southern Latin America. Eur J Wildl Res 49:112–119.

Fournier PE & Raoult D. 2003. Comparison of PCR and serology assays for an early diagnosis of acute Q fever. J Clin Microbiol. 41: 5094-5098

Frangoulidis D, Splettstoesser WD, Landt O, Dehnhardt J, Henning K, Hilbert A, Bauer

T, Antwerpen M, Meyer H, Walter MC, Knobloch JKM. 2013. Microevolution of the

Chromosomal Region of Acute Disease Antigen A (adaA) in the Query (Q) Fever Agent

Coxiella burnetii. PLoS ONE 8(1):e53440.

Fraser D. 2014. The globalisation of farm animal welfare. OIE Revue Scientifique et

Technique. 33(1): 33-38

Gad Baneth. Tick-borne infections of animals and humans: a common ground. 2014. Int

J Parasitol. 44:591–596

Gallina S. and Lopez Arevalo H. 2008. Odocoileus virginianus. The IUCN Red List of

Threatened Species 2008: e.T42394A10691422. http://dx.doi.org/10.2305/IUCN.UK.2008.RLTS.T42394A10691422.en . Downloaded on 04 November 2015.

298

Garcia-Ispierto I, Almería S, López-Gatius F. 2011. Coxiella burnetii seropositivity is highly stable throughout gestation in lactating high-producing dairy cows. Reprod

Domest Anim. 46:1067-1072

Garcia-Ispierto I, Tutusaus J, López-Gatius F. 2014. Does Coxiella burnetii affect reproduction in cattle? A clinical update. Reprod Domest Anim. 49(4): 529-535

Gardon J, Heraud JM, Laventure S, Ladam A, Capot P, Fouquet E, Favre J, Weber S,

Hommel D, Hulin A, Couratte Y, Talarmin A. 2011. Suburban Transmission of Q Fever in French Guiana: Evidence of a Wild Reservoir. J Infect Dis. 184: 278-284

Gauckler A, Kraus M. 1974. Q-Fieber bei Menschen und Tieren im Zoo Nürnberg. Verh.

Ber. Erkrg. Zootiere 16:207-212.

Georgiev M, Afonso A, Neubauer H, Needham H, Thiéry R, Rodolakis A, Roest HJ,

Stärk KD, Stegeman JA, Vellema P, van der Hoek W, More SJ. 2013. Qfever in humans and farm animals in four European countries, 1982 to 2010. Euro Surveill. 18(8): 20407

Gilbert M, Mitchell A, Bourn D, Mawdsley J, Cliton-Hadley R, Wint W. 2005. Cattle movements and bovine tuberculosis in Great Britain. Nature. 435(7041):491-496 doi:10.1038/nature03548

Giménez DF. 1965. Gram staining of Coxiella burnetii. J Bacteriol. 90: 834-835.

Giovannini A, Cancellotti FM, Turilli C, Randi E. 1988. Serological investigations for some bacterial and viral pathogens in fallow deer (Cervus dama) and wild boar (Sus scrofa) of the San Rossore Preserve, Tuscany, Italy. J Wildl Dis. 24(1): 127-132

Glazunova O, Roux V, Freylikman O, Sekeyova Z, Fournous G, Tyczka J, Tokarevich

N, Kovacava E, Marrie TJ, Raoult D. 2005. Coxiella burnetii genotyping. Emerg Infect

Dis. 11: 1211-1217

299

Gómez A, Aguirre AA. 2008. Infectious diseases and the illegal wildlife trade. Ann N Y

Acad Sci. 1149:16-19

González-Barrio D, Queirós J, Fernández-de-Mera IG, Ruiz-Fons F. 2014. Dynamics of individual exposure to Coxiella burnetii infection in a Q fever endemic red deer (Cervus elaphus) farm, abstr 28, p 71. Abstr Joint 8th Ticks and Tick-Borne Pathogens and 12th

Biennial Society for Tropical and Veterinary Medicine Conference. http://www.savetcon.co.za/TTP8/files/TTP%20STVM%20Poster%20abstracts.pdf

González-Barrio D, Velasco Ávila AL, Boadella M, Beltrán-Beck B, Barasona JA, Santos

JPV, Queirós J, García-Pérez AL, Barral M, Ruiz-Fons F. 2015a. Host and environmental factors modulate the exposure of free-ranging and farmed red deer (Cervus elaphus) to

Coxiella burnetii. Appl Environ Microbiol. 81:6223–6231

González-Barrio D, Maio E, Vieira-Pinto M, Ruiz-Fons F. 2015b. European Rabbits as

Reservoir for Coxiella burnetii. Emerg Infect Dis. 21:1055-58

González-Barrio D, Almería S, Caro MR, Salinas J, Ortíz JA, Gortázar C, Ruiz-Fons J.

2015c. Coxiella burnetii shedding by farmed red deer (Cervus elaphus). Transbound

Emerg Dis. 62:572-574

González-Barrio D, Martín-Hernando MP, Ruiz-Fons F. 2015d. Shedding patterns of endemic Eurasian wild boar (Sus scrofa) pathogens. Res Vet Sci. 102: 206-211

González-Barrio D, Fernández-de-Mera IG, Ortiz JA, Queiros J, Ruiz-Fons F. 2015e.

Long-term dynamics of Coxiella burnetii in farmed red deer (Cervus elaphus). Frontiers in Veterinary Science. Veterinary Infectious Diseases. In press

Gortázar C, Herrero J, Villafuerte R, Marco J. 2000. Historical examination of the status of large mammals in Aragon, Spain. Mammalia. 64(4): 411-422

300

Gortázar C, Acevedo A, Ruiz-Fons F, Vicente J. 2006. Disease risk andoverabundance of game species. Eur J Wildl Res. 52: 81-87

Gortázar C, Ferroglio E, Lutton CE, Acevedo P. 2010. Disease-related conflicts in mammal conservation. Wildl. Res. 37: 668-675

Gortazar C, Vicente J, Boadella M, Ballesteros C, Galindo RC, Garrido J, Aranaz A, de la Fuente J. 2011. Progress in the control of bovine tuberculosis in Spanish wildlife.

Veterinary Microbiology. 151(1-2): 170-178

Gortazar C, Reperant LA, Kuiken T, de la Fuente J, Boadella M, Martínez-Lopez B, Ruiz-

Fons F, Estrada-Peña A, Drosten C, Medley G, Ostfeld R, Peterson T, VerCauteren KC,

Menge C, Artois M, Schultsz C, Delahay R, Serra-Cobo J, Poulin R, Keck F, Aguirre A,

Henttonen H, Dobson AP, Kutz S, Lubroth J, Mysterud A. 2014a. Crossing the interspecies barrier: opening the door to zoonotic pathogens. PLoS Pathog. 10:e1004129

Gortazar C, Beltrán-Beck B, Garrido JM, Aranaz A, Sevilla IA, Boadella M,

Lyashchenko KP, Galindo RC, Montoro V, Domínguez L, Juste R, de la Fuente J. 2014.

Oral re-vaccination of Eurasian wild boar with Mycobacterium bovis BCG yields a strong protective response against challenge with a field strain. BMC Veterinary Research. 10,

Grażyna B, Jacek C, Maciej LR; Aleksander G, Tomasz M. 2012. Lyme disease: review.

Arch Med Sci. 8(6): 978-982.

Greth A, Calvez D, Vassart M, Lefèvre PC. 1992. Serological survey for bovine bacterial and viral pathogens in captive Arabian oryx (Oryx leucoryx Pallas, 1776). Rev Sci Tech

Off Int Epiz. 11(4): 1163-1168

301

Griffiths WM, Stevens DR, Archer JA, Asher GW, Littlejohn RP. 2010. Evaluation of management variables to advance conception and calving date of red deer (Cervus elaphus) in New Zealand venison production systems. Anim Reprod Sci. 118(2-4): 279-

296

Guatteo R, Beaudeau F, Berri M, Rodolakis A, Joly A, Seegers H. 2006. Shedding routes of Coxiella burnetii in dairy cows: implications for detection and control. Vet. Res. 37:

827-833.

Guatteo R, Beaudeau F, Joly A, Seegers H. 2007. Coxiella burnetii shedding by dairy cows. Vet Res. 38:849–860.

Guatteo R, Seegers H, Joly A, Beaudeau F. 2008. Prevention of Coxiella burnetii shedding in infected dairy herds using a phase I C. burnetii inactivated vaccine. Vaccine.

26:4320-4328

Gyuranecz M, Sulyok KM, Balla E, Mag T, Balázs A, Simor Z, Dénes B, Hornok S,

Bajnóczi P, Hornstra HM, Pearson T, Keim P, Dán A. 2015. Q fever epidemic in

Hungary, April to July 2013. Euro Surveill. 19(30): 20863

Gyuranecz M, Sulyok KM, Balla E, Mag T, Balázs A, Simor Z, Dénes B, Hornok S,

Bajnóczi P, Hornstra HM, Pearson T, Keim P, Dán A. 2014. Q fever epidemic in

Hungary, April to July 2013. Euro Surveill. 30

Hackstadt T, Peacock MG, Hitchcock PJ, Cole RL. 1985. Lipopolysaccharide variation in Coxiella burnetii: intrastrain heterogeneity in structure and antigenicity. Infect Immun.

48:359–365.

Hackstadt T. 1990. The role of lipopolysaccharides in the virulence of Coxiella burnetii.

Ann N Y Acad Sci. 590:27–32.

302

Hagen R, Heurich M, Kröschel M, Herdtfelder M. 2014. Synchrony in hunting bags:

Reaction on climatic and human induced changes? Sci Total Environ. 468-469: 140-146

Hamlin KL, Pac DF, Sime CA, DeSimone RM, Dusek GL. 2000. Evaluating the accuracy of ages obtained by two methods for Montana ungulates. J Wildl Manage. 64: 441-449

Hansen PJ. 2014. Current and future assisted reproductive technologies for mammalian farm animals. Adv Exp Med Biol. 752: 1-22

Hars PJ, Rossi S. 2009. Results of the surveillance of regulated contagious diseases in the

French wildlife. Bulletin de l'Academie Veterinaire de France. 162(3): 215-223

Hartung M. 2001. Mitteilungen der Länder über Coxiella burnetii-Nachweise in

Deutschland in: HARTUNG, M. (Hrsg.): Bericht über die epidemiologische Situation der

Zoonosen in Deutschland für 2000 - Übersicht über die Meldungen der Bundesländer.

BgVV-Hefte. 230-232

Hartley M & Gill E. 2010. Assessment and mitigation processes for disease risks associated with wildlife management and conservation interventions. Veterinary Record.

166(16): 487-490

Haydon DT, Cleaveland S, Taylor LH, Laurenson MK. 2002. Identifying reservoirs of infection: a conceptual and practical challenge. Emerg Infect Dis. 8: 1468-1473

Haydon DT. 2008. Cross-disciplinary demands of multihost pathogens. J Anim Ecol.

77(6): 1079-1081

Heinzen R, Hackstadt T, Samuel JE. 1999. Developmental biology of Coxiella burnetii.

Trends Microbiol. 7:149-154.

303

Heinzen R, Stiegler GL, Whiting LL, Schmitt SA, Mallavia LP, Frazier ME. 1990. Use of pulsed field gel electrophoresis to differentiate C. burnetii strains. Ann N Y Acad Sci.

590:504-513.

Hendrix LR, Samuel JE, Mallavia LP. 1991. Differentiation of Coxiella burnetii isolates by analysis of restriction-endonucleasedigested DNA separated by SDS-PAGE. J Gen

Microbiol. 137:269- 276

Henning K, Hilbert A, Wittstatt U. 2015. Seroepidemiological study on Q fever in wild boars in Berlin. Tierarztl Umsch. 70:41-42

Hermans MHA, Huijsmans CJJ, Schellekens JJA, Savelkoul PHM, Wever PC. 2011.

Coxiella burnetii DNA in goat milk after vaccination with Coxevac®. Vaccine. 29(15):

2653-2656

Herremans T, Hogema BM, Nabuurs M, Peeters M, Wegdam-Blans M, Schneeberger P,

Nijhuis C, Notermans DW, Galama J, Horrevorts A, van Loo IHM, Vlaminckx B, Zaaijer

HL, Koopmans MP, Berkhout H, Socolovschi C, Raoult D, Stenos J, Nicholson W,

Bijlmer H. 2013. Comparison of the performance of IFA, CFA, and ELISA assays for the serodiagnosis of acute Q fever by quality assessment. Diagnostic Microbiology and

Infectious Disease. 75(1): 16-21

Hernández S, Lyford-Pike V, Álvarez ME, Tomasina F. 2007. Q fever outbreak in an experimental wildlife breeding station in Uruguay. Rev Patol Trop. 36(2):129-140

Hernychova L, Toman R, Ciampor F, Hubalek M, Vackova J, Macela A, Skultety L.

Detection and identification of Coxiella burnetii based on the mass spectrometric analyses of the extracted proteins. Analytical Chemistry. 80(18): 7097-7104

304

Herrero J, Lovari S, Berducou C. 2008. Rupicapra pyrenaica. The IUCN Red List of

Threatened Species. Version 2014.2. www.iucnredlist.org. Downloaded on 04 October

2014

Herrero J & Pérez JM. 2008. Capra pyrenaica. The IUCN Red List of Threatened Species

2008: e.T3798A10085397. http://dx.doi.org/10.2305/IUCN.UK.2008.RLTS.T3798A10085397.en . Downloaded on

04 November 2015.

Hildebrandt A, Straube E, Neubauer H, Schmoock G. 2010. Coxiella burnetii and coinfections in Ixodes ricinus ticks in Central Germany. Vector Borne Zoonotic Dis.

11(8):1205-1207

Hillyard, P.D., 1996. Ticks of North-West Europe. In: Barnes, R.S.K., Crothers, J.H.

(Eds.), Synopses of the British Fauna (New Series)

Ho T, Htwe KK, Yamasaki N, Zhang GQ, Ogawa M, Yamaguchi T, Fukushi H, Hirai K.

1995. Isolation of Coxiella burnetii from dairy cattle and ticks, and some characteristics of the isolates in Japan. Microbiol Immunol. 39: 663-371.

Hoffman LC & Wiklund E. 2006. Game and venison—meat for the modern consumer.

Meat Sci. 74:197–208.

Hogerwerf L. van den Brom R, Roest HIJ, Bouma A, Vellema P, Pieterse M, Dercksen

D, Nielen M. 2011. Reduction of Coxiella burnetii prevalence by vaccination of goats and sheep, the Netherlands. Emerg Infect Dis. 17: 379-386

Hornstra HM, Priestley RA, Georgia SM, Kachur S, Birdsell DN, Hilsabeck R, Gates LT,

Samuel JE, Heinzen RA, Kersh GJ, Keim P, Massung RF, Pearson T. 2011. Rapid typing of Coxiella burnetii. PLoS ONE. 6(11): e26201

305

Hubalek Z, Juricova Z, SvobodovA S, Halouzka J. 1993. A serologic survey for some bacterial and viral zoonoses in game animals in the Czech Republic. J Wildl Dis. 29(4):

604-607

Huijsmans CJ, Schellekens JJ, Wever PC, Toman R, Savelkoul PH, Janse I, Hermans

MH. 2011. Single-nucleotide-polymorphism genotyping of Coxiella burnetii during a Q fever outbreak in the Netherlands. Appl Envirom Microbiol. 77(6): 2051-2057

Hung MN, Chou YF, Chen MJ, Hou MY, Lin PS, Lin CC, Lin LJ. 2010. Q fever outbreak in a small village, Taiwan. Jpn J Infect Dis. 63(3): 212-213

Hussein MF, Al-Khalifa IM, Aljumaah RS, Elnabi AG, Mohammed OB, Omer SA,

Macasero WV. 2012. Serological prevalence of Coxiella burnetii in captive wild ruminants in Saudi Arabia. Comp Clin Pathol. 21:33–38

Inoue K, Kabeya H, Fujita H, Makino T, Asano M, Inoue S, Inokuma H, Nogami S,

Maruyama US. 2011. Serological survey of five zoonoses, scrub typhus, Japanese spotted fever, tularemia, Lyme disease, and Q fever, in feral raccoons (Procyon lotor) in Japan.

Vector Borne Zoonotic Dis. 11: 15-19

Ioannou I, Chochlakis D, Kasinis N, Anayiotos P, Lyssandrou A, Papadopoulos B,

Tselentis Y, Psaroulaki A. 2009. Carriage of Rickettsia spp., Coxiella burnetii and

Anaplasma spp. by endemic and migratory wild birds and their ectoparasites in Cyprus.

Clin Microbiol Infect. 15:158–160

Ioannou I, Sandalakis V, Kassinis N, Chochlakis D, Papadopoulos B, Loukaides F,

Tselentis Y, Psaroulaki A. 2011. Tick-borne bacteria in mouflons and their ectoparasites in Cyprus. J Wildl Dis. 47(2): 300-306

306

Isken LD, Kraaij-Dirkzwager M, Vermeer-de Bondt PE, Rümke HC, Wijkmans C,

Opstelten W, Timen A. 2013. Implementation of a Q fever vaccination program for high- risk patients in the Netherlands. Vaccine. 31(23): 2617-2622

Jado I, Escudero R, Gil H, Jiménez-Alonso MI, Sousa R, García-Pérez AL, Rodríguez-

Vargas M, Lobo B, Anda P: Molecular method for identification of Rickettsia species in clinical and environmental samples. J Clin Microbiol 2006, 44:4572–4576

Jado I, Carranza-Rodríguez C, Barandika JF, Toledo A, García-Amil C, Serrano B,

Bolaños M, Gil H, Escudero R, García-Pérez AL, Olmeda AS, Astobiza I, Lobo B,

Rodríguez-Vargas M, Pérez-Arellano JL, López- Gatius F, Pascual-Velasco F, Cilla G,

Rodríguez NF, Anda P. 2012. Molecular method for the characterization of Coxiella burnetii from clinical and environmental samples: variability of genotypes in Spain. BMC

Microbiol. 12: 91

Jäger C, Willems H, Thiele D, Baljer G. 1998. Molecular characterization of Coxiella burnetii isolates. Epidemiol. Infect. 120: 157-164.

Jager C, Lautenschlager S, Willems H, Baljer G. 2002. Coxiella burnetii plasmid types

QpDG and QpH1 are closely related and likely identical. Vet. Microbiol. 89:161-166.

Jager C, Willems H, Thiele D, Baljer G. 1998. Molecular characterization of Coxiella burnetii isolates. Epidemiol Infect. 120: 157-164

Jang Y, Kim H, Heo I, Park Y, Kim S, Lee M, Choe N. 2011. Seroprevalence of Coxiella burnetii in cattle and farmraised deer in Korea. Afr J Microbiol Res. 5(24): 4234-4236

Jareño D, Viñuela J, Luque-Larena JJ, Arroyo L, Arroyo B, Mougeot F. 2015. Factors associated with the colonization of agricultural areas by common voles Microtus arvalis in NW Spain. Biol Invasions. 17(8): 2315-2327

307

Jiang Y, Shang H, Xu H, Zhu L, Chen W, Zhao L, Fang L. 2010. Simultaneous detection of porcine circovirus type 2, classical swine fever virus, porcine parvovirus and porcine reproductive and respiratory syndrome virus in pigs by multiplex polymerase chain reaction. Vet J. 183: 172-175

Jones RM, Hertwig S, Pitman J, Vipond R, Aspán A, Bölske G, McCaughey C, McKenna

JP, van Rotterdam BJ, de Bruin A, Ruuls R, Buijs R, Roest HJ, Sawyer J. 2011.

Interlaboratory comparison of real-time polymerase chain reaction methods to detect

Coxiella burnetii, the causative agent of Q fever. J Vet Diagn Invest. 23: 108-111

Jourdain E, Gibert P, Gauthier D, Fromont E, Jullien JM, Hars J. 2005. Sondage sur les maladies abortives chez les ongulés sauvages et domestiques en alpage. Enquête menée dans la RNCFS des Bauges. Faune Sauvage- 268: 24-32

Jurczynski K, Flugger M. 2005. A Q-fever infection (Coxiella burnetii) in marine mammals at Tierpark Hagenbeck in Hamburg—a case report. Verh ber Erkrg Zootiere.

42:184–191.

Kampschreur LM, Wegdam-Blans MCA, Thijsen SFT, Groot CAR, Schneeberger PM,

Hollander AAMJ, Schijen JHEM, Arents NLA, Oosterheert JJ, Wever PC. 2010. Acute

Q fever related in-hospital mortality in the Netherlands. Neth J Med. 68: 408-413

Karesh WB, Cook RA, Bennett EL, Newcomb J. 2005. Wildlife trade and global disease emergence. Emerg Infect Dis. 11: 1000-1002

Kersh GJ, Fitzpatrick KA, Self JS, Priestley RA, Kelly AJ, Ryan Lash R, Marsden-Haug

N, Nett RJ, Bjork A, Massung RF, Andersona AD. 2013. Presence and Persistence of

Coxiella burnetii in the environments of goat farms associated with a Q fever outbreak.

Appl Environ Microb. 79(5): 1697-1703

308

Kersh GJ, Lambourn DM, Self JS, Akmajian AM, Stanton JB, Baszler TV, Raverty SA,

Massung RF. 2010. Coxiella burnetii infection of a steller sea lion (Eumetopias jubatus)

Found in Washington State. J Clin Microbiol. 48(9):3428–3431

Kersh GJ, Lambourn DM, Raverty SA, Fitzpatrick KA, Self JS, Akmajian AM, Jeffries

SJ, Huggins J, Drew CP, Zaki SR, Massung RF. 2012. Coxiella burnetii Infection of

Marine Mammals in the Pacific Northwest, 1997–2010. J Wildl Dis. 48(1): 201-206

Keuling O, Baubet E, Duscher A, Ebert C, Fischer C, Monaco A, Podgórski T, Prevot C,

Ronnenberg K, Sodeikat G, Stier N, Thurfjell H. 2013. Mortality rates of wild boar Sus scrofa L. in central Europe. Eur J Wildl Res. 59(6): 805-814

Kirchgessner MS, Dubovi EJ, Whipps CM. 2012a. Seroepidemiology of Coxiella burnetii in wild white-tailed deer (Odocoileus virginianus) in New York, United States. Vector

Borne Zoonotic Dis. 12(11): 942-947

Kirchgessner MS, Dubovi EJ, Porter WF, Zylich NC, Whipps CM. 2012. Prevalence and spatial distribution of antibodies to bovine viral diarrhea virus and Coxiella burnetii in white-tailed deer (Odocoileus virginianus) in New York and Pennsylvania. J Zoo Wildl

Med. 43(3):466-472

Kirchgessner MS, Dubovi EJ, Whipps CM. 2013. Disease risk surface for Coxiella burnetii seroprevalence in white-tailed deer. Zoonoses Public Health. 60:457-460.

DOI:10.1111/zph.12023

Kita J, Anusz K. 1991. Serologic survey for bovine pathogens in free-ranging European bison from Poland. J Wildl Dis. 27(1): 16-20

Kittelberger R, Mars J, Wibberley G, Sting R, Henning K, Horner GW, Garnett KM,

Hannah MJ, Jenner JA, Pigott CJ, O'Keefe JS. 2009. Comparison of the Q-fever

309 complement fixation test and two commercial enzyme-linked immunosorbent assays for the detection of serum antibodies against Coxiella burnetii (Q-fever) in ruminants:

Recommendations for use of serological tests on imported animals in New Zealand. N Z

Vet J. 57: 262-268

Klaasen CHW, Nabuurs-Franssen MH, Tilburg JJHC, Hamans MAWM, Horrevorts AM.

2009. Multigenotype Q fever outbreak, the Netherlands. Emerg Infect Dis. 15: 613-614

Klee SR, Tyczka J, Ellerbrok H, Franz T, Linke S, Baljer G, Appel B. 2006. Highly sensitive real-time PCR for specific detection and quantification of Coxiella burnetii.

BMC Microbiol. 6, 2

Kukielka E, Barasona JA, Cowie CE, Drewe JA, Gortázar C, Cotarelo I, Vicente

J. 2013. Spatial and temporal interactions between livestock and wildlife in South

Central Spain assessed by camera traps. Preventive Veterinary Medicine. 112(3): 213-

221

Klyachko O, Stein BD, Grindle N, Clay K, Fuqua C. 2007. Localization and visu-alization of a Coxiella-type symbiont within the lone star tick, Amblyomma americanum. Appl

Environ Microbiol. 73: 6584-6594.

Kocianova E, Rehacek J, Lisak V. 1993. Transmission of antibodies to Chlamydia

Psittaci and Coxiella Burnetii through eggs and "crop milk" in pigeons. Eur J Epidemiol.

0392-2990: 209-212

Kosatsky T. 1984. Household outbreak of Q-fever pneumonia related to a parturient cat.

The Lancet. 2: 1447-1449.

Krauss H, Rottcher D, Weiss D, Danner K, Hubschle OJ. 1986. Wildlife as a potential source of infection in domestic animals, studies on game in Zambia. Anim Res Dev. 24:

41-58.

310

Kreizinger Z, Szeredi L, Bacsadi A, Nemes C, Sugár L, Varga T, Sulyok KM, Szigeti A,

Ács K, Tóbiás E, Borel N, Gyuranecz M. 2015. Occurrence of Coxiella burnetii and

Chlamydiales species in abortions of domestic ruminants and in wild ruminants in

Hungary, Central Europe. J Vet Diagn Invest. 27: 206-210

Kruse H, Kirkemo AM, Handeland K. 2004. Wildlife as source of zoonotic infections.

Emerging Infectious Diseases. 10(12): 2067-2072

Kuiken T, Ryser-Degiorgis MP, Gavier-Widén D, Gortázar C. 2011. Establishing a

European network for wildlife health surveillance. OIE Revue Scientifique et Technique.

30(3): 755-761

Kukielka E, Barasona JA, Cowie CE, Drewe JA, Gortazar C, Cotarelo I, Vicente J. 2013.

Spatial and temporal interactions between livestock and wildlife in South Central Spain assessed by camera traps. Prev Vet Med. 112: 213-221

Laddomada A. 2000. Incidence and control of CSF in wild boar in Europe. Vet Microbiol.

73(2-3): 121-130

Lagier JC, Edouard S, Pagnier I, Mediannikov O, Drancourt M, Raoult D. 2015. Current and past strategies for bacterial culture in clinical microbiology. Clin Microbiol Rev. 28

(1): 208-236

Lancia RA. 1994. Estimating the Number of Animals in Wildlife Populations. Research and Management Techniques for Wildlife and Habitats. Bethesda: The Wildlife Society;

1994. pp. 215–253.

Langley JM, Marrie TJ, Covert A, Waag DM, Williams JC. 1988. Poker players' pneumonia. An urban outbreak of Q fever following exposure to a parturient cat. N Engl

J Med. 319(6): 354-356

311

Lapointe JM, Gulland FM, Haines DM, Barr BC, Duignan PJ. 1999. Placentitis due to

Coxiella burnetii in a Pacific harbor seal (Phoca vitulina richardsi). J Vet Diagn Invest.

11: 541-543

Laricchiuta P, Patania T, Torina A, Vitale F, Gruppillo A, Domina F, Pennisi MG. 2006.

Arthropod-borne infections in lions (Panthera Leo) from the Fasano Safari Park. IUCN /

SSC Cat Specialist Group. Conference Proceeding

Laughlin T, Waag D, Williams J, Marrie T. 1991. Q fever: from deer to dog to man. The

Lancet. 337(16): 676-677

Lee KA, Wikelski M, Robinson WD, Robinson TR, Klasing KC. 2008. Constitutive immune defences correlate with life-history variables in tropical birds. J Anim Ecol.

77(2): 356-363

Lindsey P, Alexander R, Balme G, Midlane N, Craig J. 2012. Possible relationships between the south African captive-bred lion hunting industry and the hunting and conservation of lions elsewhere in Africa. S Afr J Wildl Res. 42(1): 11-22

Lloyd C, Mark F. Stidworthy, MA, Wernery U. 2010. Coxiella Burnetii abortion in captive dama gazelle (Gazella dama) in the United Arab Emirates. J Zoo Wildl Med.

41:83–89

Lochmiller RL & Deeremberg C. 2000. Trade-offs in evolutionary immunology: just what is the cost of immunity? Oikos. 88: 87-98

Lockhart MG, Graves SR, Banazis MJ, Fenwick SG, Stenos J. 2011. A comparison of methods for extracting DNA from Coxiella burnetii as measured by a duplex qPCR assay.

Lett Appl Microbiol. 52: 514-520

312

López-Olvera JR, Vidal D, Vicente J, Pérez M, Luján L, Gortázar C. 2009. Serological survey of selected infectious diseases in mouflon (Ovis aries musimon) from south- central Spain. Eur J Wildl Res. 55: 75-79

Lorenz H, Jager C, Willems H, Baljer G. 1998. PCR detection of Coxiella burnetii from different clinical specimens, especially bovine milk, on the basis of DNA preparation with a silica matrix. Appl Environ Microbiol. 64: 4234-4237.

Lovari S, Herrero J, Conroy J, Maran T, Giannatos G, Stubbe M, Aulagnier S, Jdeidi T,

Masseti M, Nader I, de Smet K, Cuzin F. 2008. Cervus elaphus. The IUCN Red List of

Threatened Species 2008: e.T41785A10541893. http://dx.doi.org/10.2305/IUCN.UK.2008.RLTS.T41785A10541893.en . Downloaded on 05 November 2015.

Ludt CJ, Schroeder W, Rottmann O, Kuehn R. 2004. Mitochondrial DNA phylogeography of red deer (Cervus elaphus). Mol Phylogenet Evol. 31: 1064-1083.

Lyons JA, Natusch DJD. 2013. Effects of consumer preferences for rarity on the harvest of wild populations within a species. Ecol Econ. 93: 278-283

Machado-Ferreira E, Dietrich G, Hojgaard A, Levin M, Piesman J, Zeidner NS, Soares

CAG. 2011. Coxiella Symbionts in the Cayenne Tick Amblyomma cajennense. Microb

Ecol. 62(1): 134-142

Madariaga MG, Rezai K, Trenholme GM, Weinstein RA. 2003. Q fever: A biological weapon in your backyard. Lancet Infect Dis. 3(11): 709-721

Madic J, Huber D, Lugovic B. 1993. Serologic survey for selected viral and rickettsial agents of Brown bears (Ursus arctos) in Croatia. J Wildl dis. 29(4): 572-576

313

Magar R, Larochelle R, Thibault S, Lamontagne L. 2000. Experimental transmission of porcine circovirus type 2 (PCV2) in weaned pigs: a sequential study. J Comp Pathol. 123:

258-269.

MAGRAMA. Mnisiterio de Agricultura. Alimentación y Medio Ambiente. PLAN

Nacional De Vigilancia Sanitaria En Fauna Silvestre

Maio E, Tania C, Balseiro A, Sevilla I, Romano A, Ortiz JA, Vieira-Pinto M, Garrido

JM, de la Lastra JM, Gortázar C. 2011. Paratuberculosis in European wild rabbits from

Iberian Peninsula. Res Vet Sci. 91(2): 212-218

Mallavia LP. 1991. Genetic of rickettsiae. Eur J Epidemiol. 7: 213-221

Marmion BP, Storm PA, Ayres JG, Semendric L, Mathews L, Winslow W, Turra M,

Harris RJ. 2005. Long-term persistence of Coxiella burnetii after acute primary Q fever.

QJM. 98: 7-20

Marmion BP, Shannon M, Maddocks I, Storm P, Penttila I. 1996. Protracted debility and fatigue after acute Q fever. Lancet. 347:977-978

Marreros N, Hüssy D, Albini S, Frey CF, Abril C, Vogt HR, Holzwarth N, Wirz-Dittus

S, Friess M, Engels M, Borel N, Willisch CS, Signer C, Hoelzle LE, Ryser-Degiorgis

MP. 2011. Epizootiologic investigations of selected abortive agents in free-ranging alpine ibex (Capra ibex ibex) in Switzerland. J Wildl Dis. 47(3): 530-543.

Marrie TJ, Embil J, Yates L. 1993. Seroepidemiology of Coxiella Burnetii among wildlife in Nova Scotia. Am J Trop Med Hyg. 49(5) :613-615

Marrie TJ, Schlech WF, Williams JC, Yates L. 1986. Q fever pneumonia associated with exposure to wild rabbits. Lancet. 1: 427-429.

314

Marrie TJ, Durant H, Williams JC, Mintz E, Waag DM. 1988. Exposure to parturient cats: A risk factor for acquisition of Q fever in maritime Canada. J Infect Dis. 158(1):

101-108

Marrie TJ. 1990. Q fever - a review. Can Vet J. 31: 555-563.

Marrie TJ. 1996. A dog-related outbreak of Q fever. Clin Infect Dis. 23(4): 753-755

Martin SW, Meek AH, Willeberg P: Veterinary Epidemiology. Measures of disease frequency Ames: Iowa State University Press 1987.

Massei G, Kindberg J, Licoppe A, Gacic D, Sprem N, Kamler J, Baubet E, Hohmann U,

Monaco A, Ozolin J, Cellina S, Podgórski T, Fonseca C, Markov N, Pokorny B, Rosell

C, Náhlik A. 2015. Wild boar populations up, numbers of hunters down? A review of trends and implications for Europe. Pest Manag. Sci. 71: 492-500.

Massung RF, Cutler SJ, Frangoulidis D. Molecular typing of Coxiella burnetii (Q Fever).

2012. Adv Exp Med Biol. 984: 381-396

Maurin M, Raoult D. 1999. Q fever. Clin Microbiol Rev 12:518-553

Mazeri S, Scolamacchia F, Handel IG, Morgan KL, Tanya VN, Bronsvoort BMC. 2013.

Risk factor analysis for antibodies to Brucella, Leptospira and C. burnetii among cattle in the Adamawa Region of Cameroon: a cross-sectional study. Trop Anim Health Prod.

45: 617-623

McCaul TF, Williams JC. 1981. Developmental cycle of Coxiella burnetii: structure and morphogenesis of vegetative and sporogenic differentiations. J Bacteriol. 147: 1063-

1076.

315

McCaul TF. 1991. The development cycle of Coxiella burnetii, p. 223–258. In J. C.

Williams and H. A. Thompson (ed.), Q fever: the biology of Coxiella burnetii. CRC Press,

Inc., Boca Raton, Fla.

McCulloch CE, Searle SR, Neuhaus JM. 2008. Generalized, linear, and mixed models,

2nd ed. Wiley, Hoboken, NJ.

McQuiston JH, Childs JE. 2002. Q fever in humans and animals in the United States.

Vector Borne Zoonotic Dis. 2(3): 179-191

Medić A, Dželalija B, Polićo VP, Margan IG, Turković B, Gilić V. 2005. Q fever epidemic among employees in a factory in the suburb of Zadar, Croatia. Croat Med J.

46(2): 315-319

Meerburg BG & Chantal CBEM. 2011. The role of wild rodents in spread and transmission of Coxiella burnetii needs further elucidation. Wildlife Res. 38: 617-625

Meng XJ, Lindsay DS, Sriranganathan N. 2009. Wild boars as sources for infectious diseases in livestock and humans. Philos Trans R Soc B. 364: 2697-2707

Mengeling WL. 2006. Porcine parvovirus. In: Straw, B.E., Zimmerman, J.J., D'Allaire,

S., Taylor, D.J. (Eds.), Diseases of Swine. Blackwell Publishing, Ames, pp. 373–385.

Meredith AL, Cleaveland SC, Denwood MJ, Brown JK, Shaw DJ. 2014. Coxiella burnetii

(Q-Fever) seroprevalence in prey and predators in the United Kingdom: evaluation of infection in wild rodents, foxes and domestic cats using a modified ELISA. Transbound

Emerg Dis. In press. DOI:10.1111/tbed.12211

Metcalf CJE, Birger RB, Funk S, Kouyos RD, Lloyd-Smith JO, Jansen VAA. 2015. Five challenges in evolution and infectious diseases. Epidemics. 10: 40-44

316

Mick V, Le Carrou G, Corde Y, Game Y, Jay M, Garin-Bastuji B. 2014. Brucella melitensis in France: Persistence in wildlife and probable spillover from Alpine ibex to domestic animals. PLoS ONE. 9(4): e94168

Million M, Raoult D. 2015. Recent advances in the study of Q fever epidemiology, diagnosis and management. J Infection. 71(S1): S2-S9

Million M, Thuny F, Richet H, Raoult D. 2010. Long-term outcome of Q fever endocarditis: a 26-year personal survey. Lancet Infect Dis. 10:527-535

Milner JM, Nilsen EB, Andreassen HP. 2007. Demographic side effects of selective hunting in ungulates and carnivores. Cons Biol. 21:36–47

Minor C, Kersh GJ, Gelatt T, Kondas AV, Pabilonia KL, Weller CB, Dickerson BR,

Duncan CG. 2013. Coxiella burnetii in northern fur seals and steller sea lions of Alaska.

J Wildl Dis. 49(2): 441-446

Mitchell-Jones AJ, Amori G, Bogdanomicz W, Krystufek B, Beijnders PJ, Spitzenberger

F, Stubbe M, Thissen JBM, Vohralik V, Zima J. 1999. Atlas of European mammals.

London: Academic Press; 1999.

Müller T, Freuling CM, Wysocki P, Roumiantzeff M, Freney J, Mettenleiter TC, Vos A.

2015. Terrestrial rabies control in the European Union: Historical achievements and challenges ahead. Veterinary Journal. 203(1): 10-17

Moller AP, Christe P, Eritzoe J, Mavarez J. 1998. Condition, disease and immune defence. Oikos. 83:301–306

Monnerot M, Vigne JD, Biju-Duval C, Casane D, Callou C, Hardy C, et al. 1994. Rabbit and man: genetic and historic approach. Genet Sel Evol. 26: 167-182

317

Montejo-Baranda M, Corral-Carranceja J, Aguirre-Errasti C. Q fever in the Basque

Country: 1981–1984. Rev Infect Dis 1985, 7:700–701

Montizaan MGE & Siebenga S. 2010. WBE-databank, populatie- en afschotcijfers

(Nieuwsbrief No. 2013) Koninklijke Nederlandse Jagers Vereniging.

Morroy G, Prins J, Bergevoet R, Schneeberger P, Bor HHJ, van der Hoek W, Hautvast J,

Wijkmans CJ, Peters JB, Polder JJ. 2012. Of goats and humans; the societal costs of the

Dutch Q fever saga. Int J Infect Dis. 16, pe266.

Morroy G, van Asseldonk MAPM, Bontje DM, Backer JA, Roest HIJ, Roermund HJW,

Bergevoet RHM, Prins J, Hoek WVD, Polder JJ. 2013. The societal costs of the Dutch Q fever outbreak: evaluation of past and future control strategies. In: Med–Vet–Net

Association International Scientific Conference, Lyngby, Denmark, p. 14

Morroy G, Van Der Hoek W, Albers J, Coutinho RA, Bleeker-Rovers CP, Schneeberger

PM. 2015. Population screening for chronic Q-fever seven years after a major outbreak.

2015. Plos One. 10(7): e0131777

Mostafavi E, Rastad H, Khalili M. 2012. Q fever: An emerging public health concern in

Iran. Asian J Epidemiol. 5(3): 66-73

Müller T, Hahn EC, Tottewitz F, Kramer M, Klupp BG, Mettenleiter TC, Freuling C.

2011. Pseudorabies virus in wild swine: A global perspective. Arch Virol. 156(10): 1691-

1705

Müller T, Klupp BG, Freuling C, Hoffmann B, Mojcicz M, Capua I, Palfi V, Toma B,

Lutz W, Ruiz-Fons F, Gortrzar C, Hlinak A, Schaarschmidt U, Zimmer K, Conraths FJ,

Hahn EC, Mettenleiter TC. 2010. Characterization of pseudorabies virus of wild boar origin from Europe. Epidemiol Infect. 138(11): 1590-1600

318

Muñoz PM, Boadella M, Arnal M, de Miguel MJ, Revilla M, Martínez D, Vicente J,

Acevedo P, Oleaga T, Ruiz-Fons F, Marín CM, Prieto JM, de la Fuente J, Barral M,

Barberán M, de Luco DF, Blasco JM, Gortázar C. 2010. Spatial distribution and risk factors of brucelosis in Iberian wild ungulates. BMC Infect Dis. 10:46

Myers E, Ehrhart EJ, Charles B, Spraker T, Gelatt T, Duncan C. 2013. Apoptosis in normal and Coxiella burnetii–infected placentas from Alaskan northern fur seals

(Callorhinus ursinus). Vet Pathol. 50(4): 622-625

Neagari Y, Sakai T, Nogami S, Kaiho I, Katoh C. 1998. Incidence of Antibodies in

Raccoon Dogs and Deer inhabiting Suburban Areas. Kansen Shigaku Zasshi. 72(4): 331-

334

Nelder MP, Reeves WK, Adler PH, Wozniak A, Wills W. 2009. Ectoparasites and associated pathogens of free-roaming and captive animals in zoos of South Carolina.

Vector Borne Zoonotic Dis. 9(5): 469-477

Nguyen SV, Hirai K. 1999. Differentiation of Coxiella burnetii isolates by sequence determination and PCR-restriction fragment length polymorphism analysis of isocitrate dehydrogenase gene. FEMS Microbiol Lett. 180: 249-254.

Nusinovici S, Frössling J, Widgren S, Beaudeau F, Lindberg A. 2015. Q fever infection in dairy cattle herds: Increased risk with high wind speed and low precipitation.

Epidemiol Infect. 143(15): 3316-3326

O'Connor BA, Tribe IG, Givney R. 2015. A windy day in a sheep saleyard: An outbreak of Q fever in rural South Australia. Epidemiol Infect. 143(2):391-398

Ohlson A, Malmsten J, Frössling J, Bölske G, Aspán A, Dalin AM, Lindberg A. 2014.

Surveys on Coxiella burnetii infections in Swedish cattle, sheep, goats and moose. Acta

Vet Scand. 56:39

319

Olea L, & San Miguel-Ayanz A. 2006. The Spanish dehesa. A traditional Mediterranean silvopastoral system linking production and nature conservation. Grassland Science in

Europe. 11: 3-13

Omsland A, Cockrell DC, Howe D, Fischer ER, Virtaneva K, Sturdevant DE, Porcella

SF, Heinzen RA. 2009. Host cell-free growth of the Q fever bacterium Coxiella burnetii.

Proc Natl Acad Sci U S A. 106: 4430-4434.

Oporto B, Barandika JF, Hurtado A, Aduriz G, Moreno B, Garcia-Perez AL. 2006.

Incidence of ovine abortion by Coxiella burnetii in northern Spain. Ann Y Acad Sci.

1078: 498-501.

Palomo LJ, Gisbert J, Blanco JC. 2007. Atlas y libro rojo de los mamíferos terrestres de

España. Dirección General para la Biodiversidad–SECEM– SECEMU, Madrid, Spain.

Pannwitz G, Freuling C, Denzin N, Schaarschmidt U, Nieper H, Hlinak A, Burkhardt S,

Klopries M, Dedek J, Hoffmann L, Kramer M, Selhorst T, Conraths FJ, Mettenleiter T,

Müller T. 2012. A long-termserological survey on Aujeszky's disease virus infections in wild boar in east Germany. Epidemiol Infect. 140: 348-358.

Panaiotov S, Ciccozzi M, Brankova N, Levterova V, Mitova-Tiholova M, Amicosante

M, Rezza G, Kantardjiev T. 2009. An outbreak of Q fever in Bulgaria. Annali dell'Istituto

Superiore di Sanita. 45(1): 83-86

Parker ID, Lopez RR, Karthikeyan R, Silvy NJ, Davis DS, Cathey JC. 2015. A model for assessing mammal contribution of Escherichia coli to a Texas floodplain. Wildlife Res.

42(3): 217-222.

Parker NR, Bararlet JH, Morton Bell A. 2006. Q Fever. Lancet. 367: 679-688

320

Parisi A, Fraccalvieri R, Cafiero M, Miccolupo A, Padalino I, Montagna C, Capuano F,

Sottili R. 2006. Diagnosis of Coxiella burnetii-related abortion in Italian domestic ruminants using single-tube nested PCR. Vet Microbiol. 118: 101-106

Pascual-Velasco F. 1996. Fiebre Q. Junta de Castilla-León, Consejería de Sanidad y

Bienestar Social, Zamora, España.

Pascucci I, Domenico MD, Dall’Acqua F, Sozio G, Camma C. 2015. Detection of Lyme

Disease and Q fever agents in wild rodents in Central Italy. Vector Borne Zoonotic Dis.

15(7): 404-411

Pays O, Benhamou S, Helde R, Gerard JF. 2007. The dynamics of group formation in large mammalian herbivores: an analysis in the European roe deer. Anim Behav. 74(5):

1429-1441

Pearson T, Hornstra HM, Hilsabeck R, Gates LT, Olivas SM, Birdsell DM, Hall CM,

German S, Cook JM, Seymour ML, Priestley RA, Kondas AV, Friedman CLC, RP Price,

Schupp JM, Liu CM, Price LB, Massung RF, Kersh GJ, Keim P. 2014. High prevalence and two dominant host-specific genotypes of Coxiella burnetii in U.S. milk. BMC

Microbiology. 14:41

Pejsak ZK, Truszczynski MJ. 2006. Aujeszky's disease (pseudorabies). In: Straw, B.E.,

Zimmerman, J.J., D'Allaire, S., Taylor, D.J. (Eds.), Diseases of Swine. Blackwell

Publishing, Ames, pp. 419–433.

Pérez-Gallardo F, Clavero G, Hernández S. 1952. Investigations on the epidemiology of

Q fever in Spain; mountain rabbits and dormouses as a reservoir of Coxiella burnetii. Rev

Sanid Hig Pública. 26:81-87.

321

Perugini AG, Capuano F, Esposito A, Marianelli C, Martucciello A, Iovane G, Galiero

G. 2009. Detection of Coxiella burnetii in buffaloes aborted fetuses by IS111 DNA amplification: A preliminary report. Res Vet Sci. 87: 189-191

Piñero A, Ruiz-Fons F, Hurtado A, Barandika JF, Atxaerandio R, García-Pérez AL. 2014.

Changes in the dynamics of Coxiella burnetii infection in dairy cattle: an approach to match field data with the epidemiological cycle of C. burnetii in endemic herds. J Dairy

Sci. 97: 2718-2730

Piñero A, Barandika JF, García-Pérez AL, Hurtado A. 2015. Genetic diversity and variation over time of Coxiella burnetii genotypes in dairy cattle and the farm environment. Infection, Genetics and Evolution. 31: 231-235

Pioz M, Loison A, Gauthier D, Gibert P, Jullien JM, Artois M. 2008b. Diseases and reproductive success in a wild mammal: example in the alpine chamois. Oecologia. 155:

691-704. DOI: 10.1007/s00442-007-0942-5

Pioz M, Loison A, Gibert P, Jullien JM, Artois M, Gilot-Fromont E. 2008a. Antibodies against Salmonella is associated with reduced reproductive success in female alpine chamois (Rupicapra rupicapra). Can J Zool. 86(10): 1111-1120

Pope JH, Scott W, Dwyer R. 1960. Coxiella burnetii in kangaroos and kangaroo ticks in western Queensland. Aust J Exp Biol Med Sci. 38:17-27

Potter AS, Banazis MJ, Yang R, Reid SA, Fenwick SG. 2011. Prevalence of Coxiella

Burnetii in Western grey Kangaroos (Macropus Fuliginosus) In Western Australia. J

Wildl Dis. 47(4): 821-828

322

Prokešová J, Barančeková M, Homolka M. 2006. Density of red and roe deer and their distribution in relation to different habitat characteristics in a floodplain forest. Folia Zool.

55(1): 1-14

Psaroulaki A, Hadjichristodoulou C, Loukaides F, Soteriades E, Konstantinidis A,

Papastergiou P, Ioannidou MC, Tselentis Y. 2006. Epidemiological study of Q fever in humans, ruminant animals, and ticks in Cyprus using a geographical information system.

Eur J Clin Microbiol Infect Dis. 25: 576-586.

Psaroulaki A, Chochlakis D, Ioannou I, Angelaki E, Tselentis Y. 2014a. Presence of

Coxiella burnetii in fleas in Cyprus. Vector-Borne Zoonotic Dis. 14(9): 685-687

Psaroulaki A, Chochlakis D, Angelakis E, Ioannou I, Tselentis Y. 2014b. Coxiella burnetii in wildlife and ticks in an endemic área. Trans R Soc Trop Med Hyg. 108(10):

625-631

Randolph SE, EDEN-TBD sub-project team. 2010. Human activities predominate in determining changing incidence of tick-borne encephalitis in Europe. Euro Surveill.

15(27): 24-31.

Randhawa AS, Kelly VP, Baker UEF. 1977. Agglutinins to Coxiella burnetii and

Brucella spp. with particular reference to Brucella canis, in wild animals of southern

Texas. J Am Vet Med Assoc. 171: 939-942

Ransom SE, Huebner RJ. 1951. Studies on the resistance of Coxiella burnetii to physical and chemical agents. Am J Hyg. 53: 110-119

Raoult D, Torres H, Drancourt M. 1991. Shell-vial assay: evaluation of a new technique for determining antibiotic susceptibility, tested in 13 isolates of Coxiella burnetii.

Antimicrob Agents Chemother. 35: 2070-2077

323

Raoult D, Laurent JC, Mutillod M. 1994. Monoclonal antibodies to Coxiella burnetii for antigenic detection in cell cultures and in paraffin-embedded tissues. Am J Clin Pathol.

101: 318-320.

Raoult D, Houpikian P, Dupont HT, Riss JM, Arditi-Djiane J, Brouqui P. 1999. Treatment of Q fever endocarditis: comparison of 2 regimens containing doxycycline and ofloxacin or hydroxychloroquine. Arch Intern Med. 159:167-173

Real R, Barbosa AM, Rodríguez R, García FJ, Vargas JM, Palomo LJ, Delibes M. 2009.

Conservation biogeography of ecologically interacting species: the case of the Iberian lynx and the European rabbit. Divers Distrib. 15: 390-400.

Reeves WK, Loftis AD, Sanders F, Spinks MD, Wills W, Denison AM, Dasch GA. 2006.

Borrelia, Coxiella, and Rickettsia in Carios capensis (Acari: Argasidae) from a brown pelican (Pelecanus occidentalis) rookery in South Carolina, USA. Experimental and

Applied Acarology. 39(3-4): 321-329

Rehácek J, Krauss H, Kocianová E, Kovácová E, Hinterberger G, Hanák P, Tóma V.

1993. Studies of the prevalence of Coxiella burnetii, the agent of Q fever, in the foothills of the southern Bavarian Forest, Germany. Zentralbl Bakteriol. 278: 132-138.

Rehacek J, Vosta J, Tarasevic IV, Brezina R, Jablonskaja VA, Plotnikova LF, Fetisova

NF, Hanak P. 1977. Rickettsioses studies. 3. Natural foci of rickettsioses in south

Bohemia. Bulletin of the Worl Health Organization. 55(4): 455-462

Reimer LG. 1993. Q fever. Clin Microbiol Rev. 6:193-198.

Reusken C, van der Plaats R, Opsteegh M, de Bruin A, Swart A. 2011. Coxiella burnetii

(Q fever) in Rattus norvegicus and Rattus rattus at livestock farms and urban locations in the Netherlands; could Rattus spp. represent reservoirs for (re)introduction? Prev Vet

Med. 101: 124-130

324

Riemann HP, Behymer DE, Franti CC, Crabb C, Schwab RG. 1979. Survey of Q fever agglutinins in birds and small rodents in northern California, 1975-76. J Wildl Dis. 15(4):

515-523

Rijks JM, Roest HIJ, van Tulden PW, Kik MJL, Ijzer J, Gröne A, IJzer JI. 2011. Coxiella burnetii infection in roe deer during Q fever epidemic, The Netherlands. Emerg Infect

Dis. 17: 2369-2371

Robinson SJ, Neitzel DF, Moen RA, Craft ME, Hamilton KE, Johnson LB, Mulla DJ,

Munderloh UG, Redig PT, Smith KE, Turner CL, Umber JK, Pelican KM. 2015. Disease

Risk in a Dynamic Environment: The Spread of Tick-Borne Pathogens in Minnesota,

USA. EcoHealth. 12(1): 152-163

Rodolakis A. 2006. Q fever, state of art: Epidemiology, diagnosis and prophylaxis. Small

Rum Res. 62: 121-124.

Rodolakis A, Berri M, Héchard C, Caudron C, Souriau A, Bodier CC, Blanchard B,

Camuset P, Devillechaise P, Natorp JC, Vadet JP, Arricau-Bouvery N. 2007. Comparison of Coxiella burnetii shedding in milk of dairy bovine, caprine, and ovine herds. J Dairy

Sci. 90(12): 5352-5360

Rodolakis A, Clement P, Cochonneau D, Beaudeau F, Sarradin P, Guatteo R. 2009.

Investigation of humoral and cellular immunity of dairy cattle after one or two year of vaccination with a phase I Coxiella vaccine. Procedia Vaccinol. 1: 85-88

Rodríguez-Estévez V, García A, Peña F, Gómez AG. 2009. Foraging of Iberian fattening pigs grazing natural pasture in the dehesa. Livest Sci. 120(1-2): 135-143

325

Rodríguez-Refojos C &, Zuberogoitia, I. 2011 -Sized Carnivores in Agricultural

Landscapes pp. 105-126. Middle-sized carnivores in mosaic landscapes: The case of

Biscay (SW Europe)

Roest HIJ, Tilburg JJHC, van der Hoek W, Vellema P, van Zijderveld FG, Klassen CHW,

Raoult D. 2011a. The Q fever epidemic in the Netherlands: history, onset, response and reflection. Epidemiol Infect. 139: 1-12

Roest HIJ, Ruuls RC, Tilburg JJHC, Nabuurs-Franssen MH, Klaassen CHW, Vellema P, van den Brom R, Dercksen D, Wouda W, Spierenburg MAH, van der Spek AN, Buijs R, de Boer AG, Willemsen PTJ, van Zijderveld FG. 2011b. Molecular epidemiology of

Coxiella burnetii from ruminants in Q fever outbreak, The Netherlands. Emerg Infect Dis.

17(4): 668-675

Roest HJ, van Gelderen B, Dinkla A, Frangoulidis D, van Zijderveld F, Rebel J, van

Keulen L. 2012. Q Fever in Pregnant Goats: Pathogenesis and Excretion of Coxiella burnetii. PLoS ONE. 7(11): e48949

Roest HIJ, Post J, van Gelderen B, van Zijderveld FG, Rebel JMJ. 2013a. Q fever in pregnant goats: humoral and cellular immune responses. Vet Res. 44:67

Roest HIJ, van Solt CB, Tilburg JJHC, Klaassen CH, Hovius EK, Roest FT, Vellema P, van den Brom R, van Zijderveld FG.2013b. Search for possible additional reservoirs for human Q fever, the Netherlands. Emerg Infect Dis. 19: 834-835

Roic B, Jemersic L, Terzic S, Keros T, Balatinec J, Florijancic T. 2012. Prevalence of antibodies to selected viral pathogens in wild boars (Sus scrofa) in Croatia in 2005–06 and 2009–10. J Wildl Dis. 48:131-137.

326

Romero CH, Meade PN, Shultz JE, Chung HY, Gibbs EP, Hahn EC, Lollis G. 2001.

Venereal transmission of pseudorabies viruses indigenous to feral swine. J Wildl Dis. 37:

289-296.

Rossi S, Artois M, Pontier D, Crucière C, Hars J, Barrat J, Pacholek X, Fromont E. 2005a.

Long-term monitoring of classical swine fever in wild boar (sus scrofa sp.) using serological data. Vet Res. 36(1): 27-42

Rossi S, Fromont E, Pontier D, Crucière C, Hars J, Barrat J, Pacholek X, Artois M. 2005b.

Incidence and persistence of classical swine fever in free-ranging wild boar (Sus scrofa).

Epidemiol Infect. 133(3): 559-568

Rousset E, Durand B, Berri M, Dufour P, Prigent M, Russo P, Delcroix T, Touratier A,

Rodolakis A, Aubert M. 2007. Comparative diagnostic potential of three serological tests for abortive Q fever in goat herds.Vet Microbiol. 124: 286-297

Rousset E, Durand B, Champio, JL, Prigent M, Dufour P, Forfait C, Marois M, Gasnier,

T, Duquesne V, Thiéry R, Aubert MF. 2009b. Efficiency of a phase 1 vaccine for the reduction of vaginal Coxiella burnetii shedding in a clinically affected goat herd. Clin

Microbiol Infect. 15(2):188-189

Ruiz-Fons F, Vicente J, Vidal D, Höfle U, Villanúa D, Gauss C, Segalés J, Almería S,

Montoro V, Gortázar, C. 2006a. Seroprevalence of six reproductive pathogens in

European wild boar (Sus scrofa) from Spain: The effect on wild boar female reproductive performance. Theriogenology. 65(4): 731-743

Ruiz-Fons, F., Fernández-de-Mera, I.G., Acevedo, P., Höfle, U., Vicente, J., de la Fuente,

J., Gortazár, C. 2006b. Ixodid ticks parasitizing Iberian red deer (Cervus elaphus hispanicus) and European wild boar (Sus scrofa) from Spain: Geographical and temporal

Veterinary Parasitology. Volume 140, Issue 1-2, 31 August 2006, Pages 133-142

327

Ruiz-Fons F, Vidal D, Höfle U, Vicente J, Gortázar C. 2007. Aujeszky's disease virus infection patterns in European wild boar. Vet Microbiol. 120: 241-250

Ruiz-Fons F, Reyes-García AR, Alcaide V, Gortázar C. 2008a. Spatial and temporal evolution of Bluetongue virus in wild ruminants, Spain. Emerg Infect Dis. 14: 951-953

Ruiz-Fons F, Rodríguez O, Torina A, Naranjo V, Gortázar C, de la Fuente J. 2008b.

Prevalence of Coxiella burnetii infection in wild and farmed ungulates. Vet Microbiol.

126: 282-286

Ruiz-Fons F, Segalés J, Gortázar C. 2008c. A review of viral diseases of the European wild boar: effects of population dynamics and reservoir role. Vet J. 176: 158-169

Ruiz-Fons F, Vidal D, Vicente J, Acevedo P, Fernández-de-Mera IG, Montoro V,

Gortázar C. 2008. Epidemiological risk factors of Aujeszky's disease in wild boars (Sus scrofa) and domestic pigs in Spain. Eur J Wildl Res. 54: 549-555

Ruiz-Fons F, Astobiza I, Barral M, Barandika JF, García-Pérez AL. 2010a. Modification of a commercial ELISA to detect antibodies against Coxiella burnetii in wild ungulates: application to population surveillance, abstr 6, p 16. Abstr 9th Biennial Conference,

European Wildlife Disease Association.

Ruiz-Fons F & Gilbert L. 2010b. The role of deer as vehicles to move ticks, Ixodes ricinus, between contrasting habitats. Int J Parasitol. 40: 1013-1020

Ruiz-Fons F, Astobiza I, Barandika JF, Hurtado A, Atxaerandio R, Juste RA, García-

Pérez AL. 2010c: Seroepidemiological study of Q fever in domestic ruminants in semiextensive grazing systems. BMC Vet Res. 6:3

328

Ruiz-Fons F. 2012. Coxiella burnetii infection, p 409 – 412. In Gavier-Widen D, Duff

JP, Meredith A (ed), Infectious diseases of wild mammals and birds in Europe, 1st ed.

Wiley-Blackwell, Chichester, United Kingdom.

Ruiz-Fons F, Acevedo P, Sobrino R, Vicente J, Fierro Y, Fernández-de-Mera IG. 2013.

Sex-biased differences in the effect of host individual, host population and environmental traits driving tick parasitism in red deer. Front Cell Infect Microbiol. 3:23

Ruiz-Fons F, Sánchez-Matamoros A, Gortázar C, Sánchez-Vizcaíno JM. 2014a. The role of wildlife in bluetongue virus maintenance in Europe: Lessons learned after the natural infection in Spain. Virus Research. 182: 50-58

Ruiz-Fons F, González-Barrio D, Aguilar-Ríos F, Soler AJ, Garde JJ, Gortázar C,

Fernández-Santos MR. 2014b. Infectious pathogens potentially transmitted by semen of the black variety of the manchega sheep breed: health constraints for conservation purposes. Anim Reprod Sci. 149: 152-157

Ruiz-Fons F. 2015. A review of the current status of relevant zoonotic pathogens in wild swine (Sus scrofa) populations: changes modulating the risk of transmission to humans.

Transbound Emerg Dis. In press.

Runge M, von Keyserlingk M, Braune S, Becker D, Plenge-Bonig A, Freise JF, Pelzd HJ,

Estherd A. 2013. Distribution of rodenticide resistance and zoonotic pathogens in

Norway rats in Lower Saxony and Hamburg, Germany. Pest Manag Sci. 69: 403-408

Rypuła K, Krasińska M, Kita J, Płoneczka-Janeczko K, Kapuśniak W. 2011. The prevalence of specific antibody to selected viral and bacterial infections in wild ruminants in Poland. Centr Eur J Immunol. 36(3): 180-183

329

Saegerman C, Speybroeck N, Dal Pozzo F, Czaplicki G. 2015. Clinical Indicators of

Exposure to Coxiella burnetii in Dairy Herds. Transbound Emerg Dis. 62(1): 46-54

Saegerman C, Speybroeck N, Czaplicki G. 2011, Diagnostic decision tree based on epidemiological survey. In: Proceedings of the 2nd European Ceva Q fever symposium.

Barcelona, Spain, pp 8-9.

Sáenz de Buruaga M, Lucio AJ, Purroy J. 1991. Reconocimiento de sexo y edad en especies cinegéticas. Diputación Foral de Álava, Vitoria (128 pp.)

Saito M, Koike F, Momose H, Mihira T, Uematsu S, Ohtani T, Sekiyama K. 2012.

Forecasting the range expansion of a recolonising wild boar Sus scrofa population. Wildl

Biol. 18: 383-392

Salazar DC. 2009. Distribuição e estatuto do veado e corço em Portugal. Master’s thesis.

University of Aveiro, Aveiro, Portugal.

Salinas J, Caro MR, Vicente J, Cuello F, Reyes-García AR, Buendía AJ, Rodolakis A,

Gortázar C. 2009: High prevalence of antibodies against Chlamydiaceae and

Chlamydophila abortus in wild ungulates using two “in house” blocking-ELISA tests.

Vet Microbiol. 135: 46-53

Salinas J, Ortega N, Borge C, Rangel MJ, Carbonero A, Perea A, Caro MR. 2012.

Abortion associated with Chlamydia abortus in extensively reared Iberian sows. Vet J.

194: 133-134

Salwa A, Anusz K, Arent Z, Paprocka G, Kita J. 2007. Seroprevalence of selected viral and bacterial pathogens in free-ranging European bison from the Białowieza Primeval

Forest (Poland). Pol J Vet Sci. 10(1): 19-23

330

Sambrook J, Fritsch EF, Maniatis T. 1989. Molecular cloning: a laboratory manual.

Second edition. Cold Spring Harbor Laboratory Press. New York. USA.

Samuel JE, Frazier ME, Mallavia LP. 1985. Correlation of plasmid type and disease caused by Coxiella burnetii. Infect Immun 49: 775–779

Sánchez J, Souriau A, Buendía AJ, Arricau-Bouvery N, Martínez CM, Salinas J,

Rodolakis A, Navarro JA. 2006. Experimental Coxiella burnetii infection in pregnant goats: a histopathological and immunohistochemical study. J Comp Pathol. 135: 108-115

Sánchez-Vizcaíno JM, Mur L, Gomez-Villamandos JC, Carrasco L. 2015. An update on the epidemiology and pathology of African swine fever. J Comp Pathol. 152(1): 9-21

Sanford SE, Josephson GK, MacDonald A. 1994. Coxiella burnetii (Q fever) abortion storms in goat herds after attendance at an annual fair. Can Vet J. 35:376-378.

Santiago-Moreno J, Carvajal A, Astorga RJ, Coloma MA, Toledano-Díaz A, Gómez-

Guillamon F, Salas-Vega R, López-Sebastián A. 2011. Potential impact of diseases transmissible by sperm on the establishment of Iberian ibex (Capra pyrenaica) genome resource banks. Eur J Wildl Res. 57: 211-216

Santos AS, Tilburg JJHC, Botelho A, Barahona MJ, Núncio MS, Nabuurs-Franssen MH,

Klaassen CH. 2012. Genotypic diversity of Coxiella burnetii isolates from Portugal based on MST and MLVA typing. Int J Med Microbiol. 302: 253-256

Sawyer LA, Fishbein DB, McDade JE. 1987. Q fever: current concepts. Rev Infect Dis.

9: 935-946.

Schimmer B, Luttikholt S, Hautvast JL, Graat EAM, Vellema P, van Duynhoven YTHP.

2011. Seroprevalence and risk factors of Q fever in goats on commercial dairy goat farms in the Netherlands, 2009-2010. BMC Vet Res. 7:81

331

Schleenvoigt BT, Sprague LD, Mertens K, Moog U, Schmoock G, Wolf G, Neumann M,

Pletz MW, Neubauer H. 2015. Acute Q fever infection in Thuringia, Germany, after burial of roe deer fawn cadavers (Capreolus capreolus): A case report. New Microbes

New Infect. 8: 19-20

Schöning JM, Cerny N, Prohaska S, Wittenbrink MM, Smith NH, Bloemberg G, Pewsner

M, Schiller I, Origgi FC, Ryser-Degiorgis M.P. 2013. Surveillance of bovine tuberculosis and risk estimation of a future reservoir formation in wildlife in Switzerland and

Liechtenstein. PLoS ONE. 8(1): e54253

Schröder HD. 1998. Q-fever in zoo-kept ungulates. Berl Munch Tierarztl Wochenschr.

111(5): 173-174

Schulze C, Hlinak A, Wohlsein P, Kutzer P, Müller T. 2010. Spontaneous Aujeszky's disease (pseudorabies) in European wild boars (Sus scrofa) in the federal state of

Brandenburg, Germany. Berl Munch Tierarztl Wochenschr 123: 359-364

Schulze C, Neumann G, Grütze I, Engelhardt A, Mirle C, Ehlert F, Hlinak A. 2003. Case report: porcine circovirus type 2-infection in a European wild boar (Sus scrofa) in the state of Brandenbug, Germany. Dtsch Tierarztl Wochenschr. 110: 426-428

Scott GH, Williams JC, Stephenson EH. 1987. Animal models in Q fever: pathological responses of inbred mice to phase I Coxiella burnetii. J Gen Microbiol. 133: 691-700.

Scott GH, Williams JC. 1990. Susceptibility of Coxiella burnetii to chemical disinfectants. Ann N Y Acad Sci. 590: 291-296

Segalés J, Allan GM, Domingo M. 2012. Porcine circoviruses. In: Zimmerman JJ,

Karriker LA, Ramírez A, Schwartz KJ, Stevenson GW (Eds.), Diseases of Swine.

Blackwell Publishing, Ames, pp. 405–417

332

Segalés J, Calsamiglia M, Olvera A, Sibila M, Badiella L, Domingo M. 2005.

Quantification of porcine circovirus type 2 (PCV2) DNA in serum and tonsillar, nasal, tracheobronchial, urinary and faecal swabs of pigs with and without postweaning multisystemic wasting syndrome (PMWS). Vet Microbiol. 111: 223-229

Seguin G, Lair S, Dallaire AD. 2008. Muskoxen (Ovibos moschatus) of Nunavik: state of health and food safety. Wildlife Health Centre Newsletter. 2008. 13: 6-7

Sekeyova Z, Roux V, Raoult D. 1999. Intraspecies diversity of Coxiella burnetii as revealed by com1 and mucZ sequence comparison. FEMS Microbiol Lett. 180: 61-67

Shin GW, Kim EJ, Lee HB, Cho HS. 2014. The Prevalence of Coxiella burnetii Infection in Wild Korean Water Deer, Korea. J Vet Med Sci. 76(7): 1069-1071

Shivaprasad HL, Cadenas MB, Diab SS, Nordhausen R, Bradway D, Crespo R,

Breitschwerdt EB. 2008. Coxiella-Like infection in psittacines and a toucan. Avian Dis.

52(3): 426-432

Shury TK, Nishi JS, Elkin BT, Wobeser GA. 2015. Tuberculosis and brucellosis in wood bison (Bison bison athabascae) in Northern Canada: A renewed need to develop options for future management. J Wildl Dis. 51(3): 543-554

Sidi-Boumedine K, Rousset E. 2011. Molecular epidemiology of Q fever: a review of

Coxiella burnetii genotyping methods and main achievements. Euro Reference. 5:30-37

Sidwell RW, Lundgren DL, Bushman JB, Thorpe BD. 1964. The occurrence of a possible epizootic of Q fever in fauna of the Great Salt Lake Desert of Utah. Am J Trop Med Hyg.

13(5): 754-762

Simmert J, Heckel JO, Rietschel W, Kimmig P, Sting R. 1998. Zoonotic aspects of a

Coxiella burnetii infection in farmed fallow deer (Dama dama) a case report. European

333

Association of Zoo- and Wildlife Veterinarians (EAZWV). Second scientific meeting,

May 21-24. Chester, UK.

Smith DJW, Derrick EH. 1940. Studies on the epidemiology of Q fever. I. The isolation of six strains of Rickettsia burneti from the tick Haemaphysalis humerosa. Aust J Exp

Biol Med Sci. 18: 1-5

Socolovschi C, Reynaud P, Kernif T, Raoult D, Parola P. 2012. Rickettsiae of spotted fever group, Borrelia valaisiana, and Coxiella burnetii in ticks on passerine birds and mammals from the Camargue in the south of France. Ticks Tick Borne Dis. 3: 354-359

Sprong H, Tijsse-Klasen E, Langelaar M, De Bruin A, Fonville M, Gassner F, Takken

W, Van Wieren S, Nijhof A, Jongejan J, Maassen CBM, Scholte EJ, Hovius JW, Hovius

KE, Spitalska E, Van Duynhoven YT. 2012. Prevalence of Coxiella burnetii in ticks after a large outbreak of Qfever. Zoonoses Public Health. 59: 69-75

Spyridaki I, Gikas A, Kofteridis D, Psaroulaki A, Tselentis Y. 1998. Q fever in the Greek island of Crete: detection, isolation and molecular identification of eight strains of

Coxiella burnetii from clinical samples. J Clin Microbiol. 36: 2063-2067

Spyridaki I, Psaroulaki A, Loukaides F, Antoniou M, Hadjichristodolou C, Tselentis Y.

2002. Isolation of Coxiella burnetii by a centrifugation shell-vial assay from ticks collected in Cyprus: detection by nested polymerase chain reaction (PCR) and by PCR restriction fragment length polymorphism analyses. Am J Trop Med Hyg. 66: 86-90

Stalis IH, Rideout BA. 1996. Q fever in two species of exotic ruminants. Proc Am Assoc

ZooVet. 1996: 471-473

Stein A & Raoult D. 1992. Detection of Coxiella burnetti by DNA amplification using polymerase chain reaction. J Clin Microbiol. 30: 2462-2466

334

Stein A & Raoult D. 1999. Pigeon pneumonia in provence: a bird-born q fever outbreak.

Clinical infectious diseases. 29: 617-620

Stephen SA, Rao KN. 1979. Coxiellosis in reptiles of South Kanara District, Karnataka.

Indian J Med Res. 70: 937-941

Stöbel K, Schönberg A, Staak C. 2002. A new non-species dependent ELISA for detection of antibodies to Borrelia burgdorferi s.l. in zoo animals. Int J Med Microbiol

291: 88-89

Stoenner HG, Lackman DB. 1960. The biologic properties of Coxiella burnetii isolated from rodents collected in Utah. Am J Epidemiol. 71(1): 45-51

Sutton TC & Subbarao K. 2015. Development of animal models against emerging coronaviruses: From SARS to MERS coronavirus. Virology. 479-480: 247-258

Sutmoller P. 2002. The fencing issue relative to the control of foot-and-mouth disease.

In: Gibbs EPJ, Bokma BH, editors. Domestic Animal/Wildlife Interface: Issue for

Disease Control, Conservation, Sustainable Food Production, and Emerging Diseases.

Annals of the New York Academy of Sciences. (Vol. 969), New York, NY:

New York Acad Sciences. p. 191–200

Schneider HP. 2012. The history of veterinary medicine in Namibia. J S Afr Vet Assoc.

83(1):11

Judge J, McDonald RA, Walker N, Delahay RJ. 2011. Effectiveness of biosecurity measures in preventing badger visits to farm buildings. PLoS One. 6(12):e28941

Owens M & Owens D. 1980. The fences of death. AfrWildl. 34:25-75

335

Sulyok KM, Kreizinger Z, Hornstra HM, Pearson T, Szigeti A, Dán T, Balla E, Keim PS,

Gyuranecz M. 2014. Genotyping of Coxiella burnetii from domestic ruminants and human in Hungary: Indication of various genotypes. BMC Veterinary Research. 10, 107

Sulyok KM, Hornok S, Abichu G, Erdelyi, K, Gyuranecz M. 2014. Identification of novel

Coxiella burnetii genotypes from Ethiopian ticks. PLoS ONE 9(11):e113213

Sulyok KM, Kreizinger Z, Hornstra HM, Pearson T, Szigeti A, Dán Á, Balla E, Keim PS,

Gyuranecz M. 2006. Genotyping of Coxiella burnetii from domestic ruminants and human in Hungary: indication of various genotypes. BMC Vet Res. 10: 107.

Swift L, Hunter PR, Lees AC, Bell DJ. 2007. Wildlife trade and the emergence of infectious diseases. EcoHealth. 4(1): 25-30

Svraka S, Toman R, Skultety L, Slaba K, Homan WL: Establishment of a genotyping scheme for Coxiella burnetii. FEMS Microbiol Lett 2006, 254:268–274

Szymańska-Czerwińska M, Galińska EM, Niemczuk K, Knap JP. 2015. Prevalence of

Coxiella burnetii infection in humans occupationally exposed to animals in Poland.

Vector-Borne Zoonotic Dis. 15(4): 261-267

Swift L, Hunter PR, Lees AC, Bell DJ. 2007. Wildlife trade and the emergence of infectious diseases. EcoHealth. 4(1): 25-30

Tatsumi N, Baumgartner A, Qiao Y, Yamamoto I, Yamaguchi K. 2006. Detection of

Coxiella burnetii in chicken market eggs and mayonnaise. Ann N Y Acad Sci. 1078: 502-

505

Tauscher K, Pietschmann J, Wernike K, Teifke JP, Beer M, Blome S. 2015. On the situation of African swine fever and the biological characterization of recent virus isolates. Berl Munch Tierarztl Wochenschr. 128(5-6): 169-176

336

Tellez A, Sainz C, Echevarria C, de Carlos S, Fernandez MV, Leon P, Brezina R. 1988.

Q fever in Spain: acute and chronic cases, 1981-1985. Reviews of Infectious Diseases.

10(1):198-202

Tempelman C, Prins J, Koopmans C. 2011. Economic Consequences of the Q-Fever

Outbreak. SEO Economisch Onderzoek, Amsterdam.

Thiele D, Willems H, Köpf G, Krauss H. 1993. Polymorphism in DNA restriction patterns of Coxiella burnetii isolates investigated by pulsed field gel electrophoresis and image analysis. Eur J Epidemiol. 9:419-425

Thompson M, Mykytczuk N, Gooderham K, Schulte-Hostedde A. 2012. Prevalence of the bacterium Coxiella burnetii in wild rodents from a Canadian natural environment park. Zoonoses Public Health. 59: 553-560

Thornton PK. 2010. Livestock production: recent trends, future prospects. Philos Trans

R Soc B. 365(1554): 2853-2867

Tilburg JJ, Rossen JW, van Hannen EJ, Melchers WJ, Hermans MH, van de Bovenkamp

J, Roest HJ, de Bruin A, Nabuurs-Franssen MH, Horrevorts AM, Klaassen CH. 2012b.

Genotype diversity of Coxiella burnetii in the 2007-2010 Q fever outbreak episodes in

The Netherlands. J Clin Microbiol. 50: 1076-1078.

Tilburg JJHC, Melchers WJ, Petterson AM, Rossen JM, Hermans MH, van Hannen EJ,

Nabuurs-Franssen MH, de Vries MC, Horrevorts AM, Klaassen CHW. 2010.

Interlaboratory evaluation of different extraction and real-time PCR methods for detection of Coxiella burnetii DNA in serum. J Clin Microbiol. 48: 3923-3927

Tilburg JJHC, Roest HIJ, Buffet S, Nabuurs- Franssen MH, Horrevorts AM, Raoult D,

Klaassen KW. 2012a: Epidemic genotype of Coxiella burnetii among goats, sheep, and humans in the Netherlands. Emerg Infect Dis. 18: 887-888

337

Tilburg JJHC, Roest HJIJ, Nabuurs-Franssen MH, Horrevorts AM, Klaassen CHW. 2012.

Genotyping reveals the presence of a predominant genotype of Coxiella burnetii in consumer milk products. J Clin Microbiol. 50: 2156-2158

Tilburg JJHC. Molecular investigation of the Q fever epidemic in the Netherlands. The largest outbreak caused by Coxiella burnetii ever reported. 2013. PhD. Thesis, Radboud

University Nijmegen.

Tissot-Dupont H, Amadei MA, Nezri M, Raoult D. 2004. Wind in November, Q fever in

December. Emerg Infect Dis. 10: 1264-1269

Tissot-Dupont H, Raoult D. 2008. Q fever. Infect Dis Clin N Am. 22: 505-514

Thompson M, Mykytczuk N, Gooderham K, Schulte-Hostedde A. 2012. Prevalence of the bacterium Coxiella burnetii in wild rodents from a canadian natural environment park.

Zoonoses and Public Health. 59(8): 553-560

To H, Hotta A, Zhang GQ, Nguyen SV, Ogawa M, Yamaguchi T, Fukushi H, Amano K,

Hirai K. 1998. Antigenic characteristics of polypeptides of Coxiella burnetii isolates.

Microbiol Immunol. 42: 81-85

To H, Kako N, Zhang GQ, Otsuka H, Ogawa M, Ochiai O, Nguyen SV, Yamaguchi T,

Fukushi H, Nagaoka N, Akiyama M, Amano K, Hirai. 1996. Q fever pneumonia in children in Japan. J Clin Microbiol. 34: 647-651.

To H, Sakai R, Shirota K, Kano C, Abe S, Sugimoto T, Takehara K, Morita C, Takashima

I, Maruyama T, Yamaguchi T, Fukushl H, Hirai K. 1995. Coxiellosis in domestic and wild birds from Japan. Journal of wildlife diseases. 34(2): 310-316

Toledo A, Jado I, Olmeda AS, Casado-Nistal MA, Gil H, Escudero R, Anda P. 2009.

Detection of Coxiella burnetii in Ticks Collected from Central Spain. Vector Borne

Zoonotic Dis. 9(5): 465-8

338

Toma T, Mancini F, Di Luca M, Cecere JG, Bianchi R, Khoury C, Quarchioni E, Manzia

F, Rezza G, Ciervo A. 2014. Detection of microbial agents in ticks collected from migratory birds in Central Italy. Vector-borne and zoonotic diseases. 14(3): 199-205

Torina A, Naranjo V, Pennisi MG, Patania T, Vitale F, Laricchiuta P, Alongi A, Scimeca

S, Kocan KM, de la Fuente J. 2007. Serologic and molecular characterization of tick- borne pathogens in lions (Panthera Leo) from the Fasano Safari Park, Italy. J Zoo Wildl

Med. 38(4): 591-593

Tozer SJ, Lambert SB, Strong CL, Field HE, Sloots TP, Nissen MD. 2014. Potential animal and environmental sources of Q fever infection for humans in Queensland.

Zoonoses Public Health. 61: 105-112.

Tutusaus J, López-Gatius F, Almería S, Serrano B, Monleón E, Badiola J, García-Ispierto

I. 2013. No detectable precolostral antibody response in calves born from cows with cotyledons positive for Coxiella burnetii by quantitative PCR. Acta Vet Hung. 61:4432-

4441

Tylewska-Wierzbanowska S, Rumin W, Lewkowicz H, Sikorski S. 1991. Epidemic of Q fever in Leszno District in Poland. Eur J Epiderniol. 0392-2990: 307-309

Van Asseldonk MAPM, Prins J, Bergevoet RHM. 2013. Economic assessment of Q fever in the Netherlands. Preventive Veterinary Medicine. 112(1-2): 27-34

Van Asseldonk MAPM, Bontje DM, Backer JA, van Roermund HJW, Bergevoet RHM.

2015. Economic aspects of Q fever control in dairy goats. Preventive Veterinary

Medicine. 121(1-2): 115-122

Van den Brom R, Vellema P. 2009. Q fever outbreaks in small ruminants and people in the Netherlands. Small Rumin Res. 86(1-3): 74-79

339

Van den Brom R, van Engelen E, Roest HIJ, van der Hoek W, Vellema P. 2015. Coxiella burnetii infections in sheep or goats: an opinionated review. Vet Microbiol. In press.

DOI:10.1016/j.vetmic.2015.07.011

Van Der Hoek W, Morroy G, Renders NHM, Wever PC, Hermans MHA, Leenders

ACAP, Schneeberger PM. 2012. Epidemic Q fever in humans in the Netherlands. Adv

Exp Med Biol. 984: 329-364

Van Heerden J, Mills MG, Van Vuuren MJ, Kelly PJ, Dreyer MJ. 1995. An investigation into the health status and diseases of wild dogs (Lycaon pictus) in the Kruger National

Park. J S Afr Vet Assoc. 66(1): 18-27

Van Schaik EJ, Chen C, Mertens K, Weber MM, Samuel JE. 2013. Molecular pathogenesis of the obligate intracellular bacterium Coxiella burnetii. Nature Reviews

Microbiology. 11(8): 561-573

Vander Wal E, Paquet PC, Messier F, McLoughlin PD. 2013. Effects of phenology and sex on social proximity in a gregarious ungulate. Can J Zool. 91: 601-609

Vellema P, van den Brom R. 2014. The rise and control of the 2007-2012 human Q fever outbreaks in the Netherlands. Small Rumin Res. 118(1-3): 69-78

Verin R, Varuzza P, Mazzei M, Poli A. 2014. Serologic, molecular, and pathologic survey of pseudorabies virus infection in hunted wild boars (Sus scrofa) in Italy. J Wildl Dis. 50:

559-565

Viana M, Mancy R, Biek R, Cleaveland S, Cross PC, Lloyd-Smith JO, Haydon DT. 2014.

Assembling evidence for identifying reservoirs of infection. Trends Ecol Evol 29: 270-

279.

340

Vicente J, Höfle U, Garrido JM, Fernández-De-Mera IG, Juste R, Barral M, Gortazar C.

2006. Wild boar and red deer display high prevalences of tuberculosis-like lesions in

Spain. Vet Res. 37: 107-119

Vicente J, Ruiz-Fons F, Vidal D, Höfle U, Acevedo P, Villanúa D, Fernández-de-Mera

IG, Martín MP, Gortázar C. 2005. Serosurvey of Aujeszky's disease virus infection in

European wild boar in Spain. Vet Re. 156: 408-412

Vicente J, Segalés J, Höfle U, Balasch M, Plana-Durán J, Domingo M, Gortázar C, 2004.

Epidemiological study on porcine circovirus type 2 (PCV2) infection in the European wild boar (Sus scrofa). Vet Re. 35: 243-253.

Víchová B, Majláthová V, Nováková M, Stanko M, Hviscová I, Pangrácová L,

Chrudimský T, Curlík J, Petko B. 2014. Anaplasma infections in ticks and reservoir host from Slovakia. Infect Genet Evol. 22: 265-272

Walander P. 2012. Six years of estimating roe and fallow deer density with distance sampling at the Koberg estate. Bachelor in Wildlife Ecology. Swedish University of

Agricultural Sciences. Faculty of Natural Resources and Agricultural Sciences.

Department of Ecology. Grimsö Wildlife Research Station

Webster JP, Lloyd G, Macdonald DW. 1995. Q fever (Coxiella burnetii) reservoir in wild brown rat (Rattus norvegicus) populations inthe UK. Parasitology. 110: 31-35

White CL, Schuler KL, Thomas NJ, Webb JL, Saliki JT, Ip HS, Dubey JP, Frame ER.

2013. Pathogen exposure and blood chemistry in the Washington, USA, population of northern sea otters (Enhydra lutris kenyoni). J Wildlife Dis. 49(4): 887-899

341

Whitney EAS, Massung RF, Candee AJ, Ailes EC, Myers LM, Patterson NE, Berkelman

RL. 2009. Seroepidemiologic and occupational risk survey for Coxiella burnetii antibodies among US veterinarians. Clin Infect Dis. 48: 550-557

Widmer CE, Azevedo FCC, Almeida AP, Ferreira F, Labruna MB. 2011. Tick-borne bacteria in free-living jaguars (Panthera onca) in Pantanal, Brazil. Vector-Borne

Zoonotic Dis. 11(8): 1001-1005

Willeberg P, Ruppanner R, Behymer DE, Haghighi S, Kaneko JJ, Franti CE. 1980.

Environmental exposure to Coxiella burnetii: A sero-epidemiologic survey among domestic animals. Am J Epidemiol. 111(4): 437-443

Willems H, Thiele D, Frolich-Ritter R, Krauss H. 1994. Detection of Coxiella burnetii in cow's milk using the polymerase chain reaction (PCR). Zentralbl Veterinarmed. 41: 580-

587

Wisser J, Strauss G, Henschke J. 1993. Chronisch verflaufende Q-Fieber-Infektion beim

Moschusochsen (Ovibos moschatus). Visz. 35: 181-187.

Wobeser GA. 1994. Investigation and management of disease in wild animals. Plenum,

New York, NY.

Woc-Colburn AM, Garner MM, Bradway D, West G, D'Agostino J, Trupkiewicz J, Barr

B, Anderson SE, Rurangirwa FR, Nordhausen RW. 2008. Fatal Coxiellosis in Swainson's

Blue Mountain Rainbow Lorikeets (Trichoglossus haematodus moluccanus). Vet Pathol.

45(2): 247-254

Wood JLN, Leach M, Waldman L, MacGregor H, Fooks AR, Jones KE, Restif O,

Dechmann D, Hayman DTS, Baker KS, Peel AJ, Kamins AO, Fahr J, Ntiamoa-Baidu Y,

Suu-Ire R, Breiman RF, Epstein JH, Field HE, Cunningham AA. 2012. A framework for the study of zoonotic disease emergence and its drivers: Spillover of bat pathogens as a

342 case study. Philosophical Transactions of the Royal Society B: Biological Sciences.

367(1604): 2881-2892

Woldehiwet Z. 2004. Q fever (coxiellosis): epidemiology and pathogenesis. Res Vet Sci.

77: 93-100

Woodbury MR, Berezowski J, Haigh J. 2006. An estimation of reproductive performance of farmed elk (Cervus elaphus) in North America. Can Vet J. 47: 60-64

Yadav MP, Sethi MS. 1979. Poikilotherms as reservoirs of Q-fever (Coxiella burnetii) in

Uttar Pradesh. J Wildl Dis. 15(1): 15-17

Yadav MP, Sethi MS. 1980. A study on the reservoir status of Q-fever in avifauna, wild mammals and poikilotherms in Uttar Pradesh (India). Int J Zoonoses. 7(2): 85-89

Zachos FE & Hartl GB. 2011. Phylogeography, population genetics and conservation of the European red deer Cervus elaphus. Mammal Rev. 41: 138-150

Zanet S, Trisciuoglio A, Bottero E, Fernandez De Mera IG, Gortázar C, Carpignano MG,

Ferroglio E. 2014. Piroplasmosis in wildlife: Babesia and Theileria affecting free-ranging ungulates and carnivores in the Italian Alps. Parasite Vector. 7:70

Zarnke RL. 1983. Serologic survey for selected microbial pathogens in Alaskan wildlife.

J Wildl Dis. 19(4):324-329

Zhang GQ, Nguyen SV, To H, Ogawa M, Hotta A, Yamaguchi T, Kim HJ, Fukushi H,

Hirai K. 1998. Clinical evaluation of a new PCR assay for detection of Coxiella burnetii in human serum samples. J Clin Microbiol. 36: 77-80

Zhang G, To H, Russell KE, Hendrix LR, Yamaguchi T, Fukushi H, Hirai K, Samuel JE.

2005. Identification and characterization of an immunodominant 28-kilodalton Coxiella

343 burnetii outer membrane protein specific to isolates associated with acute disease. Infect

Immun. 73: 1561-1567.

Saegerman C, Speybroeck N, Dal Pozzo F, Czaplicki G. 2015. Clinical Indicators of

Exposure to Coxiella burnetii in Dairy Herds. Transbound Emerg Dis. 62(1): 46-54

Ruiz-Fons F, Astobiza I, Barandika JF, Juste RA, Hurtado A, García-Pérez AL. 2011.

Measuring antibody levels in bulk-tank milk as an epidemiological tool to search for the status of Coxiella burnetii in dairy sheep. Epidemiol Infect. 139(10): 1631-1636

Eibach R, Bothe F, Runge M, Ganter M. 2013. Long-term monitoring of a Coxiella burnetii-infected sheep flock after vaccination and antibiotic treatment under field conditions. Berl Munch Tierarztl Wochenschr. 126(1-2): 3-9

Hamann HP, Volmer R, Wimmershof N, Ballmann G, Zschöck M. 2009. Q-fever- vaccination in sheep. Tierarztl Umsch. 64(4): 188-190

Brooks DL, Ermel RW, Franti CE, Ruppanner R, Behymer DE, Williams JC, Stephenson

EH, Stephenson JC. 1986. Q fever vaccination of sheep: challenge of immunity in ewes.

Am J Vet Res. 47(6): 1235-1238

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